Thiamine: Catalytic Mechanisms in Normal and Disease States

Catalytic mPthanirmr in normal and Dirtare httr e d i t e d b y Frank Jordan Rutgers University Newark, New Jersey, U...

Author: Frank Jordan | Mulchand S. Patel

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Catalytic mPthanirmr in normal and Dirtare httr e d i t e d b y

Frank Jordan Rutgers University Newark, New Jersey, U.S.A.

mulchand S. Pate1 School of Medicine and Biomedical Sciences State University of New York at BufSalo Bufsalo, New York, U.S.A.

m M A R C E L

D E K K E R

MARCELDEKKER, INC.

NEWYORK BASEL

Although great care has been taken to provide accurate and current information, neither the author(s) nor the publisher, nor anyone else associated with this publication, shall be liable for any loss, damage, or liability directly or indirectly caused or alleged to be caused by this book. The material contained herein is not intended to provide specific advice or recommendations for any specific situation. 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 A catalog record for this book is available from the Library of Congress. ISBN: 0-8247-4062-9 This book is printed on acid-free paper. Headquarters Marcel Dekker, Inc., 270 Madison Avenue, New York, NY 10016, U.S.A. tel: 212-696-9000; fax: 212-685-4540 Distribution and Customer Service Marcel Dekker, Inc., Cimarron Road, Monticello, New York 12701, U.S.A. tel: 800-228-1160; fax: 845-796-1772 Eastern Hemisphere Distribution Marcel Dekker AG, Hutgasse 4, Postfach 812, CH-4001 Basel, Switzerland tel: 41-61-260-6300; fax: 41-61-260-6333 World Wide Web http://www.dekker.com The publisher offers discounts on this book when ordered in bulk quantities. For more information, write to Special Sales/Professional Marketing at the headquarters address above. Copyright n 2004 by Marcel Dekker, Inc. All Rights Reserved. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage and retrieval system, without permission in writing from the publisher. Current printing (last digit): 10 9 8 7 6 5 4 3 2 1 PRINTED IN THE UNITED STATES OF AMERICA

OXIDATIVE STRESS AND DISEASE Series Editors

LESTERPACKER,PH.D. ENRIQUE CADENAS, M.D., PH.D. University of Southern California School of Pharmacy Los Angeles, California

1. Oxidative Stress in Cancer, AIDS, and Neurodegenerativa Diseases, edited by Luc Montagnier, Rene Olivier, and Catherine Paslquier 2. Understanding the Process of Aging: The Roles of Mitochondria, Free Radicals, and Antioxidants, edited by Enrique Cadenas and Lester Packer 3. Redox Regulation of Cell Signaling and Its Clinical Application, edited by Lester Packer and Junji Yodoi 4. Antioxidants in Diabetes Management, edited by Lester Packer, Peter Rosen, Hans J. Tritschler, George L. King, and Angelo Azzr 5. Free Radicals in Brain Pathophysiology, edited by Giuseppe Poli, Enrique Cadenas, and Lester Packer 6. Nutraceuticals in Health and Disease Prevention, edited by Klaus Kramer, Peter-Paul Hoppe, and Lester Packer 7. Environmental Stressors in Health and Disease, edited by Jurgen Fuchs and Lester Packer 8. Handbook of Antioxidants: Second Edition, Revised and Expanded, edited by Enrique Cadenas and Lester Packer 9. Flavonoids in Health and Disease: Second Edition, Revised and Expanded, edited by Catherine A. Rice-Evans and Lester Packer 10. Redox-Genome Interactions in Health and Disease, edited by Jurgen Fuchs, Maurizio Podda, and Lester Packer 11. Thiamine: Catalytic Mechanisms in Normal and Disease States, edited by Frank Jordan and Mulchand S. Pate1

Related Volumes

Vitamin E in Health and Disease: Biochemistry and Clinical Applications, edited by Lester Packer and Jurgen Fuchs Vitamin A in Health and Disease, edited by Rune Blornhoff

Free Radicals and Oxidation Phenomena in Biological Systems, edited by Marcel Roberfroid and Pedro Buc Calderon Biothiols in Health and Disease, edited by Lester Packer and Enrique Cadenas Handbook of Antioxidants, edited by Enrique Cadenas and Lester Packer Handbook of Synthetic Antioxidants, edifed by Lester Packer and Enrique Cadenas Vitamin C in Health and Disease, edited by Lester Packer and Jurgen Fuchs Lipoic Acid in Health and Disease, edited by Jurgen Fuchs, Lester Packer, and Guido Zimmer

Additional Volumes in Preparation Phytochemicals in Health and Disease, edited by Yongping Bao and Roger Fenwick Carotenoids in Health and Disease, edited by Norman 1. Krinsky, Susan T. Mayne, and Helmut Sies Herbal Medicine, edited by Lester Packer, Choon Nam Ong, and Balrry Halliwell

Series Introduction

Oxygen is a dangerous friend. Overwhelming evidence indicates that oxidative stress can lead to cell and tissue injury. However, the same free radicals that are generated during oxidative stress are produced during normal metabolism and thus are involved in both human health and disease. Free radicals are molecules with an odd number of electrons. The odd, or unpaired, electron is highly reactive as it seeks to pair with another free electron. Free radicals are generated during oxidative metabolism and energy production in the body. Free radicals are involved in: Enzyme-catalyzed reactions Electron transport in mitochondria Signal transduction and gene expression Activation of nuclear transcription factors Oxidative damage to molecules, cells, and tissues Antimicrobial action of neutrophils and macrophages Aging and disease Normal metabolism is dependent on oxygen, a free radical. Through evolution, oxygen emerged as the terminal electron acceptor for respiration. The two unpaired electrons of oxygen spin in the same direction; thus, oxygen is a biradical, but is not a very dangerous free radical. Other oxygen-derived iii

iv

Series Introduction

free radical species, such as superoxide or hydroxyl radicals, formed during metabolism or by ionizing radiation are stronger oxidants and are therefore more dangerous. In addition to research on the biological effects of these reactive oxygen species, research on reactive nitrogen species has been gathering momentum. NO, or nitrogen monoxide (nitric oxide), is a free radical generated by NO synthase (NOS). This enzyme modulates physiological responses such as vasodilation or signaling in the brain. However, during inflammation, synthesis of NOS (iNOS) is induced. This iNOS can result in the overproduction of NO, causing damage. More worrisome, however, is the fact that excess NO can react with superoxide to produce the very toxic product peroxynitrite. Oxidation of lipids, proteins, and DNA can result, thereby increasing the likelihood of tissue injury. Both reactive oxygen and nitrogen species are involved in normal cell regulation in which oxidants and redox status are important in signal transduction. Oxidative stress is increasingly seen as a major upstream component in the signaling cascade involved in inflammatory responses, stimulating adhesion molecule and chemoattractant production. Hydrogen peroxide, which breaks down to produce hydroxyl radicals, can also activate NF-nB, a transcription factor involved in stimulating inflammatory responses. Excess production of these reactive species is toxic, exerting cytostatic effects, causing membrane damage, and activating pathways of cell death (apoptosis and/or necrosis). Virtually all diseases thus far examined involve free radicals. In most cases, free radicals are secondary to the disease process, but in some instances free radicals are causal. Thus, there is a delicate balance between oxidants and antioxidants in health and disease. Their proper balance is essential for ensuring healthy aging. The term oxidative stress indicates that the antioxidant status of cells and tissues is altered by exposure to oxidants. The redox status is thus dependent on the degree to which a cell’s components are in the oxidized state. In general, the reducing environment inside cells helps to prevent oxidative damage. In this reducing environment, disulfide bonds (S—S) do not spontaneously form because sulfhydryl groups kept in the reduced state (SH) prevent protein misfolding or aggregation. This reducing environment is maintained by oxidative metabolism and by the action of antioxidant enzymes and substances, such as glutathione, thioredoxin, vitamins E and C, and enzymes such as superoxide dismutase (SOD), catalase, and the selenium-dependent glutathione and thioredoxin hydroperoxidases, which serve to remove reactive oxygen species. Changes in the redox status and depletion of antioxidants occur during oxidative stress. The thiol redox status is a useful index of oxidative stress

Series Introduction

v

mainly because metabolism and NADPH-dependent enzymes maintain cell glutathione (GSH) almost completely in its reduced state. Oxidized glutathione (glutathione disulfide, GSSG) accumulates under conditions of oxidant exposure, and this changes the ratio of oxidized to reduced glutathione; an increased ratio indicates oxidative stress. Many tissues contain large amounts of glutathione, 2–4 mM in erythrocytes or neural tissues and up to 8 mM in hepatic tissues. Reactive oxygen and nitrogen species can directly react with glutathione to lower the levels of this substance, the cell’s primary preventative antioxidant. Current hypotheses favor the idea that lowering oxidative stress can have a clinical benefit. Free radicals can be overproduced or the natural antioxidant system defenses weakened, first resulting in oxidative stress, and then leading to oxidative injury and disease. Examples of this process include heart disease and cancer. Oxidation of human low-density lipoproteins is considered the first step in the progression and eventual development of atherosclerosis, leading to cardiovascular disease. Oxidative DNA damage initiates carcinogenesis. Compelling support for the involvement of free radicals in disease development comes from epidemiological studies showing that an enhanced antioxidant status is associated with reduced risk of several diseases. Vitamin E and prevention of cardiovascular disease is a notable example. Elevated antioxidant status is also associated with decreased incidence of cataracts and cancer, and some recent reports have suggested an inverse correlation between antioxidant status and occurrence of rheumatoid arthritis and diabetes mellitus. Indeed, the number of indications in which antioxidants may be useful in the prevention and/or the treatment of disease is increasing. Oxidative stress, rather than being the primary cause of disease, is more often a secondary complication in many disorders. Oxidative stress diseases include inflammatory bowel diseases, retinal ischemia, cardiovascular disease and restenosis, AIDS, ARDS, and neurodegenerative diseases such as stroke, Parkinson’s disease, and Alzheimer’s disease. Such indications may prove amenable to antioxidant treatment because there is a clear involvement of oxidative injury in these disorders. In this series of books, the importance of oxidative stress in diseases associated with organ systems of the body is highlighted by exploring the scientific evidence and the medical applications of this knowledge. The series also highlights the major natural antioxidant enzymes and antioxidant substances such as vitamins E, A, and C, flavonoids, polyphenols, carotenoids, lipoic acid, and other nutrients present in food and beverages. Oxidative stress is an underlying factor in health and disease. More and more evidence indicates that a proper balance between oxidants and antioxidants is involved in maintaining health and longevity and that altering this

vi

Series Introduction

balance in favor of oxidants may result in pathological responses causing functional disorders and disease. This series is intended for researchers in the basic biomedical sciences and clinicians. The potential for healthy aging and disease prevention necessitates gaining further knowledge about how oxidants and antioxidants affect biological systems. Frank Jordan and Mulchand S. Patel should be congratulated for editing Thiamine: Catalytic Mechanisms in Normal and Disease States, which represents a important contribution to the Oxidative Stress and Disease Series. Thiamine is an essential component of a-keto acid dehydrogenase, mitochondrial multienzyme complexes that are important for the regulation of oxidative energy production and intermediary metabolism. The specific role that thiamine plays in pyruvate decarboxylase—the first enzyme in the pyruvate dehydrogenase complex—has been extensively investigated to reveal new structural and functional characteristics. Not surprisingly, defects in the function of thiamine-containing enzymes are associated with numerous metabolic disorders and disease states. Lester Packer Enrique Cadenas

Preface

Thiamine: Catalytic Mechanisms in Normal and Disease States brings together recent developments in thiamine diphosphate (TDP)–requiring enzymes, covering a broad field in biochemistry. TDP serves as a cofactor of various enzymes concerned mostly with decarboxylation of a-ketoacids. The mechanisms of TDP-catalyzed reactions have been extensively investigated over the past two decades, covering aspects of chemistry and biochemistry. These findings were richly covered by the proceedings of four meetings on this topic as well as two volumes specifically covering the multienzyme a-ketoacid dehydrogenase complexes (see Bibliography). These volumes provided comprehensive reviews of most recent developments in this field at the time of their publication. In May 2002, an international conference, ‘‘Thiamin, Its Biochemistry and Structural Biology,’’ was held at Rutgers University in Newark, New Jersey. This meeting prompted us to put together a volume on this theme—not as the proceedings of the conference, but rather as a compendium of important developments—by inviting selected experts to summarize their most exciting findings in the past few years. In developing this book, emphasis was placed on two specific aspects of TDP-requiring enzymes: detailing the mechanisms of catalysis and structure– function relationships, and pathophysiological aspects of a spectrum of diseases associated with TDP-requiring enzymes. There have been explosive developments on crystal structures of several TDP-dependent enzymes in recent years and these aspects are covered in depth in several chapters. Equally exciting developments in genetic defects and neuropathological findings on vii

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Preface

neurodegenerative diseases involving TDP-dependent enzymes are also covered in this volume. We thank all the contributors for their prompt, authoritative, and insightful reviews. Their combined efforts have made this volume an important resource for researchers, both newly initiated and established, in this field. We would also like to thank Dr. Lioubov G. Korotchkina and Dr. Natalia S. Nemeria for their generous assistance in realizing this volume. Frank Jordan Mulchand S. Patel

BIBLIOGRAPHY Bisswanger H, Schellenberger A, eds. Biochemistry and Physiology of Thiamin Diphosphate Enzymes. Prien, Germany: Wissenschaftlicher Verlag, 1996. Bisswanger H, Ulrich J, eds. Biochemistry and Physiology of Thiamin Diphosphate Enzymes. Weinheim, Germany: VCH, 1991. Patel MS, Roche TE, Harris RA, eds. Alpha-Keto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996. Roche TE, Patel MS, eds. Alpha-Keto Acid Dehydrogenase Complexes: Organization, Regulation, and Biomedical Ramifications. Ann. NY Acad. Sci. 573, 1989. Sable HZ, Gubler CJ, eds. Thiamin: Twenty Years of Progress. Ann. NY Acad. Sci. 378, 1982. Schellenberger A, organizer. Biochimica et Biophysica Acta, 1385, 1998. Schellenberger A, Schowen RL, eds. Thiamine Pyrophosphate Biochemistry. Boca Raton, Florida: CRC Press, 1988.

Contents

Series Introduction Preface Contributors Part I: 1.

3.

4.

iii vii xv

Introduction

Chemical Intermediates in Catalysis by Thiamine Diphosphate Perry A. Frey

Part II: 2.

Lester Packer and Enrique Cadenas

1

Biosynthesis of Thiamine and Its Phosphorylated Terms

Mechanistic and Structural Studies on Thiamine Biosynthetic Enzymes Tadhg P. Begley and Steven E. Ealick Studies on the Structure and Function of Thiamine Pyrophosphokinase Jing-Yuan Liu, David E. Timm, Robert A. Harris, and Thomas D. Hurley New Perspectives on the Cellular Role of Thiamine Triphosphate and Thiamine Triphosphatase Lucien Bettendorff and Pierre Wins

15

29

43

ix

x

Contents

Part III: 5.

6.

7.

8.

9.

10.

11.

Enzymology of Thiamine Diphosphate Enzymes

How Thiamine Works in Enzymes: Time-Resolved NMR Snapshots of TDP-Dependent Enzymes in Action Kai Tittmann, Ralph Golbik, Kathrin Uhlemann, Ludmila Khailova, Mulchand S. Patel, Frank Jordan, David M. Chipman, Ronald G. Duggleby, Gerhard Hu¨bner, and Gunter Schneider Thiamine-Dependent Enzymes as Catalysts of C–C Bond-Forming Reactions: The Role of ‘‘Orphan’’ Enzymes Michael Mu¨ller and Georg A. Sprenger Ligand-Induced Conformational Changes in Thiamine Diphosphate–Dependent Enzymes: Comparison Between Crystal and Solution Structures Stephan Ko¨nig, Michael Spinka, Erik Fiedler, Georg Wille, Johanna Brauer, Michel H. J. Koch, and Dmitri I. Svergun Enantioselective Synthesis of Hydroxy Ketones via Benzoylformate Decarboxylase- and Benzaldehyde Lyase-Catalyzed C–C Bond Formation Bettina Lingen, Martina Pohl, Ayhan S. Demir, Andreas Liese, and Michael Mu¨ller Benzoylformate Decarboxylase: Lessons in Enzymology Michael J. McLeish, George L. Kenyon, Elena S. Polovnikova, Asim K. Bera, Natalie L. Anderson, and Miriam S. Hasson New Concept on the Nature of the Induced Absorption Band of Holotransketolase Marina V. Kovina, Irina A. Sevostyanova, Olga N. Solovjeva, Ludmilla E. Meshalkina, and German A. Kochetov Structure of the a-Carbanion/Enamine Reaction Intermediate in the Active Site of Transketolase, Determined by Kinetic Crystallography Tatyana Sandalova, Erik Fiedler, Stina Thorell, Ralph Golbik, Stephan Ko¨nig, and Gunter Schneider

57

77

93

113

131

143

159

Contents

12.

13.

14.

xi

Yeast Pyruvate Decarboxylase: New Features of the Structure and Mechanism Frank Jordan, Min Liu, Eduard Sergienko, Zhen Zhang, Andrew Brunskill, Palaniappa Arjunan, and William Furey Solvent and Carbon Kinetic Isotope Effects on Active-Site and Regulatory-Site Variants of Yeast Pyruvate Decarboxylase Wen Wei, Min Liu, Lan Chen, W. Phillip Huskey, and Frank Jordan Insights into the Mechanism and Regulation of Bacterial Acetohydroxyacid Synthases David M. Chipman, Ze’ev Barak, Stanislav Engel, Sharon Mendel, and Maria Vyazmensky

173

217

233

15.

Structure and Properties of Acetohydroxyacid Synthase Ronald G. Duggleby, Luke W. Guddat, and Siew Siew Pang

16.

Exploring the Substrate Specificity of Benzoylformate Decarboxylase, Pyruvate Decarboxylase, and Benzaldehyde Lyase Petra Siegert, Martina Pohl, Malea M. Kneen, Irina D. Pogozheva, George L. Kenyon, and Michael J. McLeish

275

Benzoylformate Decarboxylase: Intermediates, Transition States, and Diversions Ronald Kluger, Qingyan Hu, and Ian F. Moore

291

17.

Part IV: 18.

251

Structure and Function of Thiamine Diphosphate Multienzyme Complexes

Structural and Functional Organization of Pyruvate Dehydrogenase Complexes Z. Hong Zhou, Lester Reed, and James K. Stoops

19.

The Pyruvate Dehydrogenase Multienzyme Complex Richard N. Perham, Jacqueline S. Milne, and Sriram Subramaniam

20.

Activation and Transfer of Lipoic Acid in Protein Lipoylation in Mammals Kazuko Fujiwara, Kazuko Okamura-Ikeda, and Yutaro Motokawa

309 331

343

xii

21.

22.

23.

24.

25.

Contents

Central Organization of Mammalian Pyruvate Dehydrogenase (PD) Complex and Lipoyl Domain–Mediated Activated Function and Control of PD Kinases and Phosphatase 1 Thomas E. Roche, Yasuaki Hiromasa, Ali Turkan, Xiaoming Gong, Tao Peng, Xiaohua Yan, Shane A. Kasten, Haiying Bao, and Jianchun Dong Physiological Effects of Replacing the PDH Complex of E. coli by Genetically Engineered Variants or by Pyruvate Oxidase John R. Guest, Ahmed M. Abdel-Hamid, Graham A. Auger, Louise Cunningham, Robin A. Henderson, Rosane S. Machado, and Margaret M. Attwood Structure and Intersubunit Information Transfer in the E. coli Pyruvate Dehydrogenase Multienzyme Complex William Furey, Palaniappa Arjunan, Andrew Brunskill, K. Chandrasekhar, Natalia S. Nemeria, Wen Wei, Yan Yan, Sheng Zhang, and Frank Jordan Structure, Function, and Regulation of Pyruvate Dehydrogenase Kinase Kirill M. Popov, Alina Tuganova, Mellissa M. BowkerKinley, Boli Hung, Pengfei Wu, C. Nicklaus Steussy, Jean Hamilton, and Robert A. Harris Three-Dimensional Structures for Components and Domain of the Mammalian Branched-Chain a-Ketoacid Dehydrogenase Complex David T. Chuang, R. Max Wynn, and Jacinta L. Chuang

Part V: 26.

27.

28.

363

387

407

433

449

Biomedical Aspects of Thiamine Diphosphate– Dependent Enzymes

Variability of Human Pyruvate Dehydrogenase Complex Deficiency Douglas S. Kerr and Christine L. Schmotzer

471

Kinetic Studies of Human Pyruvate Dehydrogenase and Its Mutants: Interactions with Thiamine Pyrophosphate Mulchand S. Patel and Lioubov G. Korotchkina

485

The Complexity of Single-Gene Disorders: Lessons from Maple Syrup Urine Disease and Thiamine Responsiveness Dean J. Danner, Eric A. Muller, and Andrea Kasinski

509

Contents

29.

30.

31.

Thiamine Pyrophosphate: An Essential Cofactor in the Mammalian Metabolism of 3-Methyl-Branched Fatty Acids Minne Casteels, Veerle Foulon, Guy P. Mannaerts, and Paul P. Van Veldhoven

525

Pathogenesis of Selective Neuronal Loss in Wernicke– Korsakoff Syndrome: Role of Oxidative Stress Paul Desjardins and Roger F. Butterworth

539

Thiamine-Responsive Megaloblastic Anemia Syndrome: Clinical Aspects and Molecular Genetics Kimihiko Oishi, George A. Diaz, and Bruce D. Gelb

549

Part VI: 32.

xiii

Concluding Remarks

Accomplishments and Future Directions Frank Jordan and Mulchand S. Patel

Index

565

569

Contributors

Ahmed M. Abdel-Hamid, B.Sc., M.Sc., Ph.D.** Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, England Natalie L. Anderson Department of Biological Sciences, Purdue University– West Lafayette, West Lafayette, Indiana, U.S.A. Palaniappa Arjunan Biocrystallography Laboratory, Veterans Affairs Medical Center, and Department of Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, U.S.A. Margaret M. Attwood, B.Sc., Ph.D. Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, England Graham A. Auger, B.Sc., Ph.D.y Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, England Haiying Bao Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A.

* Current affiliation: Department of Botany, Minia University, Minia, Egypt. y Current affiliation: The Medical School, University of Sheffield, Sheffield, England.

xv

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Contributors

Ze’ev Barak Department of Life Sciences, Ben-Gurion University of the Negev, Beer-Sheva, Israel Tadhg P. Begley, Ph.D. Department of Chemistry and Chemical Biology, Cornell University, Ithaca, New York, U.S.A. Asim K. Bera Department of Biological Sciences, Purdue University–West Lafayette, West Lafayette, Indiana, U.S.A. Lucien Bettendorff, Ph.D. Center for Cellular and Molecular Neurobiology, University of Lie`ge, Lie`ge, Belgium Mellissa M. Bowker-Kinley Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. Johanna Brauer Department of Biochemistry and Biotechnology, Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Andrew Brunskill Biocrystallography Laboratory, Veterans Affairs Medical Center, and Department of Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, U.S.A. Roger F. Butterworth, Ph.D., D.Sc. Neuroscience Research Unit, CHUM (Hoˆpital St-Luc), University of Montreal, Montreal, Quebec, Canada Minne Casteels, M.D., Ph.D. Afdeling Farmacologie, Departement Moleculaire Celbiologie, Faculty of Medicine, Katholieke Universiteit Leuven, Leuven, Belgium K. Chandrasekhar Biocrystallography Laboratory, Veterans Affairs Medical Center, and Department of Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, U.S.A. Lan Chen Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. David M. Chipman, Ph.D. Department of Life Sciences, Ben-Gurion University of the Negev, Beer-Sheva, Israel

Contributors

xvii

David T. Chuang, Ph.D. Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas, U.S.A. Jacinta L. Chuang Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas, U.S.A. Louise Cunningham, B.Sc., Ph.D. Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, England Dean J. Danner, Ph.D. Department of Human Genetics, Emory University School of Medicine, Atlanta, Georgia, U.S.A. Ayhan S. Demir Department of Chemistry, Middle East Technical University, Ankara, Turkey Paul Desjardins Neuroscience Research Unit, CHUM (Hoˆpital St-Luc), University of Montreal, Montreal, Quebec, Canada George A. Diaz, M.D., Ph.D. Departments of Pediatrics and Human Genetics, Mount Sinai School of Medicine, New York, New York, U.S.A. Jianchun Dong Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. Ronald G. Duggleby, Ph.D. Department of Biochemistry and Molecular Biology, The University of Queensland, Brisbane, Australia Steven E. Ealick, Ph.D. Department of Chemistry and Chemical Biology, Cornell University, Ithaca, New York, U.S.A. Stanislav Engel, M.Sc. Department of Life Sciences, Ben-Gurion University of the Negev, Beer-Sheva, Israel Erik Fiedler, Dr.rer.nat.** Department of Biochemistry and Biotechnology, Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Veerle Foulon, Ph.D. Afdeling Farmacologie, Departement Moleculaire Celbiologie, Faculty of Medicine, Katholieke Universiteit Leuven, Leuven, Belgium * Current affiliation: Project Manager Affilines, Scil Proteins GmbH, Halle/Saale, Germany.

xviii

Contributors

Perry A. Frey, Ph.D. Department of Biochemistry, University of Wisconsin–Madison, Madison, Wisconsin, U.S.A. Kazuko Fujiwara Institute for Enzyme Research, University of Tokushima, Tokushima, Japan William Furey, Ph.D. Biocrystallography Laboratory, Veterans Affairs Medical Center, and Department of Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, U.S.A. Bruce D. Gelb, M.D. Departments of Pediatrics and Human Genetics, Mount Sinai School of Medicine, New York, New York, U.S.A. Ralph Golbik Department of Biochemistry and Biotechnology, Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Xiaoming Gong Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. Luke W. Guddat Department of Biochemistry and Molecular Biology, The University of Queensland, Brisbane, Australia John R. Guest, B.Sc., D.Phil., F.R.S. Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, England Jean Hamilton Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. Robert A. Harris, Ph.D. Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. Miriam S. Hasson, Ph.D. Department of Biological Sciences, Purdue University–West Lafayette, West Lafayette, Indiana, U.S.A. Robin A. Henderson, B.Sc., Ph.D.** Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, England

* Current affiliation: Regulatory Affairs, Biotechnology Group, Kendle International Ltd., Ely, England.

Contributors

xix

Yasuaki Hiromasa Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. Qingyan Hu Davenport Chemical Research Laboratory, Department of Chemistry, University of Toronto, Toronto, Ontario, Canada Gerhard Hu¨bner Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Boli Hung Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. Thomas D. Hurley, Ph.D. Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. W. Phillip Huskey Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Frank Jordan, Ph.D. Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Andrea Kasinski Department of Human Genetics, Emory University School of Medicine, Atlanta, Georgia, U.S.A. Shane A. Kasten Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. George L. Kenyon The College of Pharmacy, University of Michigan, Ann Arbor, Michigan, U.S.A. Douglas S. Kerr, M.D., Ph.D. Center for Inherited Disorders of Energy Metabolism, Rainbow Babies and Children’s Hospital, and Departments of Pediatrics, Biochemistry, and Nutrition, Case Western Reserve University School of Medicine, Cleveland, Ohio, U.S.A. Ludmilla Khailova A. N. Bakh Institute of Biochemistry, Russian Academy of Sciences, Moscow, Russia Ronald Kluger, Ph.D. Davenport Chemical Research Laboratory, Department of Chemistry, University of Toronto, Toronto, Ontario, Canada

xx

Contributors

Malea M. Kneen The College of Pharmacy, University of Michigan, Ann Arbor, Michigan, U.S.A. Michel H. J. Koch, Dr.rer.nat. NCS Group, European Molecular Biology Laboratory, Hamburg Outstation, Germany German A. Kochetov, Ph.D., Dr.Sc. A. N. Belozersky Institute of PhysicoChemical Biology, Moscow State University, Moscow, Russia Stephan Ko¨nig, Dr.rer.nat. Department of Biochemistry and Biotechnology, Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Lioubov G. Korotchkina, Ph.D. Department of Biochemistry, School of Medicine and Biomedical Sciences, State University of New York at Buffalo, Buffalo, New York, U.S.A. Marina V. Kovina, Ph.D. A. N. Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow, Russia Andreas Liese Institut fu¨r Biotechnologie 2, Forschungszentrum Ju¨lich GmbH (Research Centre Ju¨lich), Ju¨lich, Germany Bettina Lingen Institute for Enzyme Technology, Heinrich-Heine-Universita¨t Du¨sseldorf, Ju¨lich, Germany Jing-Yuan Liu Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. Min Liu Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Rosane S. Machado, B.Sc., Ph.D.** Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, England Guy P. Mannaerts, M.D., Ph.D. Afdeling Farmacologie, Departement Moleculaire Celbiologie, Faculty of Medicine, Katholieke Universiteit Leuven, Leuven, Belgium

* Current affiliation: Laboratory of Molecular Medicine and Biotechnology, Centro de Biotecnologia, Porto Alegre-RS, Brazil.

Contributors

xxi

Michael J. McLeish, Ph.D. The College of Pharmacy, University of Michigan, Ann Arbor, Michigan, U.S.A. Sharon Mendel, Ph.D.** Department of Life Sciences, Ben-Gurion University of the Negev, Beer-Sheva, Israel Ludmilla E. Meshalkina, Ph.D. A. N. Belozersky Institute of PhysicoChemical Biology, Moscow State University, Moscow, Russia Jacqueline S. Milne, Ph.D. Laboratory of Biochemistry, National Cancer Institute, National Institutes of Health, Bethesda, Maryland, U.S.A. Ian F. Moorey Davenport Chemical Research Laboratory, Department of Chemistry, University of Toronto, Toronto, Ontario, Canada Yutaro Motokawa Institute for Enzyme Research, University of Tokushima, Tokushima, Japan Eric A. Muller Department of Human Genetics, Graduate Program in Genetics and Molecular Biology, Medical Scientist Training Program, Emory University School of Medicine, Atlanta, Georgia, U.S.A. Michael Mu¨ller Institut fu¨r Biotechnologie 2, Forschungszentrum Ju¨lich GmbH (Research Centre Ju¨lich), Ju¨lich, Germany Natalia S. Nemeria Department of Chemistry and the Program in Cellular and Molecular Biodynamics, Rutgers University, Newark, New Jersey, U.S.A. Kimihiko Oishi, M.D. Departments of Pediatrics and Human Genetics, Mount Sinai School of Medicine, New York, New York, U.S.A. Kazuko Okamura-Ikeda Institute for Enzyme Research, University of Tokushima, Tokushima, Japan

* Current affiliation: Department of Chemistry, University of Warwick, Coventry, England. y Current affiliation: Antimicrobial Research Centre, Department of Biochemistry, McMaster University, Hamilton, Ontario, Canada.

xxii

Contributors

Siew Siew Pang Department of Biochemistry and Molecular Biology, The University of Queensland, Brisbane, Australia Mulchand S. Patel, Ph.D. Department of Biochemistry, School of Medicine and Biomedical Sciences, State University of New York at Buffalo, Buffalo, New York, U.S.A. Tao Peng Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. Richard N. Perham, Sc.D., F.R.S. Cambridge Centre for Molecular Recognition, Department of Biochemistry, Cambridge University, Cambridge, England Irina D. Pogozheva The College of Pharmacy, University of Michigan, Ann Arbor, Michigan, U.S.A. Martina Pohl Institute for Enzyme Technology, Heinrich-Heine-Universita¨t Du¨sseldorf, Ju¨lich, Germany Elena S. Polovnikova** Department of Biological Sciences, Purdue University–West Lafayette, West Lafayette, Indiana, U.S.A. Kirill M. Popov Division of Molecular Biology and Biochemistry, School of Biological Sciences, University of Missouri–Kansas City, Kansas City, Missouri, U.S.A. Lester J. Reed, Ph.D. Department of Chemistry and Biochemistry, University of Texas at Austin, Austin, Texas, U.S.A. Thomas E. Roche, Ph.D. Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. Tatyana Sandalova Division of Molecular Structural Biology, Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm, Sweden Christine L. Schmotzer Case Western Reserve University School of Medicine, Cleveland, Ohio, U.S.A.

* Current affiliation: Kilpatrick Stockton LLP, Atlanta, Georgia, U.S.A.

Contributors

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Gunter Schneider Division of Molecular Structural Biology, Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm, Sweden Eduard Sergienko Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Irina A. Sevostyanova, Ph.D. A. N. Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow, Russia Petra Siegert Institute for Enzyme Technology, Heinrich-Heine-Universita¨t Du¨sseldorf, Ju¨lich, Germany Olga N. Solovjeva, Ph.D. A. N. Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow, Russia Michael Spinka, B.Sc. Department of Biochemistry and Biotechnology, Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Georg A. Sprenger** Institut fu¨r Biotechnologie 2, Forschungszentrum Ju¨lich GmbH (Research Centre Ju¨lich), Ju¨lich, Germany C. Nicklaus Steussy Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. James K. Stoops, Ph.D. Department of Pathology, University of Texas Health Science Center at Houston Medical School, Houston, Texas, U.S.A. Sriram Subramaniam, Ph.D. Laboratory of Biochemistry, National Cancer Institute, National Institutes of Health, Bethesda, Maryland, U.S.A. Dmitri I. Svergun, PD Dr.rer.nat.habil. NCS Group, European Molecular Biology Laboratory, Hamburg Outstation, Germany, and Institute of Crystallography, Russian Academy of Sciences, Moscow, Russia

* Current affiliation: Institut fu¨r Mikrobiologie, Universita¨t Stuttgart, Stuttgart, Germany.

xxiv

Contributors

Stina Thorell Division of Molecular Structural Biology, Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm, Sweden David E. Timm Division of Discovery Chemistry Research, Eli Lilly and Company, Indianapolis, Indiana, U.S.A. Kai Tittmann Institute of Biochemistry, Martin-Luther-Universita¨t HalleWittenberg, Halle/Saale, Germany Alina Tuganova Division of Molecular Biology and Biochemistry, School of Biological Sciences, University of Missouri–Kansas City, Kansas City, Missouri, U.S.A. Ali Turkan Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. Kathrin Uhlemann Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Paul P. Van Veldhoven, Ph.D. Afdeling Farmacologie, Departement Moleculaire Celbiologie, Faculty of Medicine, Katholieke Universiteit Leuven, Leuven, Belgium Maria Vyazmensky, Ph.D. Department of Life Sciences, Ben-Gurion University of the Negev, Beer-Sheva, Israel Wen Wei Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Georg Wille, B.Sc. Department of Biochemistry and Biotechnology, Institute of Biochemistry, Martin-Luther-Universita¨t Halle-Wittenberg, Halle/ Saale, Germany Pierre Wins Center for Cellular and Molecular Neurobiology, University of Lie`ge, Lie`ge, Belgium Pengfei Wu Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. R. Max Wynn Department of Internal Medicine, University of Texas Southwestern Medical Center, Dallas, Texas, U.S.A.

Contributors

xxv

Xiaohua Yan Department of Biochemistry, Kansas State University, Manhattan, Kansas, U.S.A. Yan Yan Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Sheng Zhang Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Zhen Zhang Department of Chemistry, Rutgers University, Newark, New Jersey, U.S.A. Z. Hong Zhou, Ph.D. Department of Pathology and Laboratory Medicine, University of Texas Health Science Center at Houston Medical School, Houston, Texas, U.S.A.

1 Chemical Intermediates in Catalysis by Thiamine Diphosphate Perry A. Frey University of Wisconsin–Madison, Madison, Wisconsin, U.S.A.

I. INTRODUCTION One of the most chemically interesting and satisfying biological mechanisms to be unveiled within the past 50 years is that of the role of thiamine diphosphate (TDP) in enzymatic catalysis. TDP serves as the indispensable coenzyme in enzymatic cleavages of carbon–carbon bonds in a,h-dicarbonyl compounds and a-hydroxycarbonyl compounds. This function of TDP is the biological raison d’eˆtre for vitamin B1, thiamine. TDP facilitates these cleavages in the active sites of many enzymes, including pyruvate decarboxylase, a-ketoacid dehydrogenase complexes, transketolase, pyruvate oxidase, acetolactase synthase, and pyruvate oxidoreductases, among others. In each case, the role of TDP is interesting and very highly satisfying in terms of chemical principles. The key to the chemical mechanism of action of TDP was revealed in 1957 by R. Breslow in the first major bio-organic chemical application of high-resolution NMR (1). Breslow discovered that 3,4-dimethylthiazolium salts underwent rapid hydrogen exchange of C2(H) with deuterium in neutral D2O, with a half-time of about 20 min in a process catalyzed by deu1

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teroxide ion (DO). This exchange implicated the ionization described by Eq. (1):

Facile ionization of C2(H) in neutral solution to form the zwitterionic, ylide-like carbanion immediately revealed the likely secret to the chemistry of TDP reactions. The transiently formed TDP-carbanion might undergo nucleophilic addition to the carbonyl groups of substrates at the active sites of enzymes. The resultant adducts possessed just the chemical properties required in the carbon–carbon bond cleavages for which the TDP enzymes had become known. The clear and logical mechanisms of decarboxylation and ketol cleavage that are so well known today were revealed, initially by the observation of the exchange of Eq. (1). The chemical role of the thiazolium ring of TDP as an electron sink is clearly defined in the mechanism of the reaction of pyruvate decarboxylase. This aspect of the participation of TDP appears in outline in Figure 1, where the quaternary nitrogen in the thiazolium ring serves as the repository of the

Figure 1 Role of TDP in the mechanism of the reaction of pyruvate decarboxylase.

Chemical Intermediates in Catalysis by TDP

3

electron pair arising from the heterolytic cleavage of the carbon dioxide from lactyl-TDP, the adduct of pyruvate with the thiazolium-ylide of TDP. The resulting hydroxyethylidene-TDP accepts a proton to form hydroxyethylTDP, the adduct of acetaldehyde with TDP. Decomposition of this adduct produces acetaldehyde and regenerates TDP. A key intermediate in the decarboxylation of pyruvate is hydroxyethylidene-TDP, widely known as the enamine intermediate. This and the closely related dihydroxyethylidene-TDP play central roles in essentially all TDPdependent reactions, as illustrated in Figure 2. The enamines can react by

Figure 2 Central roles of enamine- and acyl-TDP intermediates in thiamine-dependent reactions.

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Frey

protonation to form hydroxyethyl- or dihydroxyethyl-TDP and thence the aldehyde or ketone products. Alternatively, they can undergo oxidation to the acyl-TDPs, which can react with acyl group acceptors to form products at the oxidation level of carboxylic acids. There is a tendency to accept familiar mechanisms as if they were always obvious and as if the original experiments unmasking them were routine laboratory exercises. This is almost never true, certainly not in the field of TDP mechanisms. Prior to the unexpected discovery of the ionization of the thiazolium ring (1), many alternative chemical mechanisms for the action of TDP were under serious consideration. These mechanisms included central roles for the pseudobase of the thiazolium ring, the methylene group bridging the thiazolium and pyrimidine rings, and the 4V-amino group of the pyrimidine ring (2). The alternative mechanisms fell out of favor upon Breslow’s discovery of reaction 1 and the identification of hydroxyethyl- and dihydroxyethyl-TDP in enzymatic reactions. II. ALDEHYDE ADDITION COMPOUNDS WITH TDP The mechanisms of TDP reactions were verified and supported by the isolation and characterization of compounds from enzymatic reactions that were predicted by the potential chemical reactivity of the TDP-carbanion. These compounds included 2-(1V-)hydroxyethyl-TDP (hydroxyethyl-TDP) and 2-(1V,2V-dihydroxyethyl)-TDP (dihydroxyethyl-TDP), which proved to be true intermediates in the reactions of pyruvate decarboxylase and transketolase, respectively (3–5):

The ionized enamine forms of these compounds were found to be intermediates in many other TDP-dependent reactions, including those of the pyruvate dehydrogenase complex and pyruvate oxidase. III. ACYL-TDPs AS INTERMEDIATES Evidence for other TDP intermediates came to light early in the research on the chemical mechanisms. It seemed obvious that oxidized forms of hy-


5

droxyethyl-TDP must be transiently formed in several TDP-dependent reactions, notably pyruvate oxidase and phosphoketolase and possibly the pyruvate dehydrogenase complex. The formation of acetate or acetyl CoA as enzymatic products implied the intermediate formation of 2-acetyl-TDP (acetyl-TDP). The chemical properties of synthetic chemical models for acetyl-TDP, such as 2-acetyl-3,5-dimethylthiazolium salts, were compatible with and supported the participation of acetyl-TDP as an intermediate in enzymatic reactions (6,7). The characterization of acetyl-TDP as a compound and enzymatic intermediate lagged behind other progress in TDP research because of perceptions that it would be difficult to synthesize and would in any case be too unstable to purify and characterize. These concerns were well founded, based on the expectation that acetyl-TDP would undergo hydrolysis rapidly in neutral solutions at rates comparable to that for 2-acetyl-3,4-dimethylthiazolium ion (7). A. Implication of Acyl-TDPs in the Reactions of A-Ketoacid Dehydrogenase Complexes Evidence for the possible participation of acyl-TDPs in the reactions of Aketoacid dehydrogenase complexes inspired the eventual synthesis of acetylTDP. The complexes catalyze the overall transformations of A-ketoacids into NADH, acyl CoAs, and carbon dioxide according to Eq. (2):

The steps in the overall reactions of a-ketoacid dehydrogenase complexes in Figure 3 provide a framework for defining the issue of the involvement of acyl-TDPs in these reactions. The complexes all consist of three central enzymes, the A-ketoacid dehydrogenase (E1), the dihydrolipoyl transacylase (E2), and the dihydrolipoyl dehydrogenase (E3) (8). The steps in which each enzyme carries out its catalytic function are reversible and may be described and defined by Eq. (3a) through (3f ) in Figure 3. The putative intermediate acyl-TDP in equations (3b) and (3c) is bracketed to highlight its position in the overall mechanism. Because Eqs. (3a) through (3f ) are reversible, the intermediates should be generated by addition of the products to the enzyme complex as well as by the addition of the substrates. A high-energy intermediate, such as an acylTDP, would not be likely to accumulate in a high percentage of the active sites in either case. However, to probe the mechanism for the formation of acylTDPs, it is possible to take advantage of a well-known chemical property of

6

Frey

Figure 3 Steps in the mechanisms of the reactions of a-ketoacid dehydrogenase complexes. The mechanism of the reaction of a-ketoacid dehydrogenase complexes are represented in six steps that take place at the active sites of three enzymes, the a-ketoacid dehydrogenase, E1, the dihydrolipoyl transacylase, E2, and the dihydrolipoyl dehydrogenase, E3. One putative intermediate, an acyl-TDP, arises from dehydrogenation of the enamine-TDP intermediate at the active site of E1 in Eq. (3b), and the acyl group is transferred to the dihydrolipoyl moiety of E2 in Eq. (3c). The steps are all reversible. The first step, Eq. (3), can be made to be irreversible by excluding CO2. In that case, reversal of the reaction upon addition of NADH and the acyl CoA (RCOSCoA) can allow the accumulation of the putative intermediate acylTDP, but only when TDP is present in the active site of E1. Accumulated acyl-TDP can undergo hydrolysis in a side reaction characteristic of this compound in solution. The observation of TDP-dependent hydrolysis of cognate acyl CoAs by the aketoacid dehydrogenases supports the intermediate formation of acyl-TDPs at the active sites of these enzymes. (From Refs. 9 and 10.)

these compounds, hydrolysis to the carboxylic acid and TDP. In an effective method for probing the mechanism for the participation of an acyl-TDP, the A-ketoacid dehydrogenase complex can be incubated with NADH and the cognate acyl CoA in the absence of CO2 and tested for the hydrolysis of the acyl CoA. The absence of CO2 makes Eq. (3a) irreversible, and this should allow the putative acyl-TDP to accumulate as a dead-end species. Under these conditions, in which the acyl-TDP could not be carboxylated, its hydrolysis could take place as an alternative process, and this would lead to the observation of enzyme-catalyzed hydrolysis of the acyl CoA. To be a signal for the intermediate formation of an acyl-TDP, the mechanism of acyl CoA hydrolysis must proceed through all of the steps in the overall mechanism, except for the first step, which is blocked by the absence of CO2. The other steps can take place whenever the acyl CoA, NADH, and TDP are present. TDP would be required because purified A-ketoacid dehydrogenase


7

complexes do not have endogenous TDP and require the addition of TDP to display activity. The complexes contain the FAD required for the action of E3 and do not require added FAD to display maximum activity. The pyruvate dehydrogenase complex and A-ketoglutarate dehydrogenase complex from Escherichia coli catalyze the hydrolysis of acetyl CoA and succinyl CoA, respectively, in the absence of added CO2 in reactions that require the presence of both NADH and TDP (9,10). The simplest and most obvious rationale is that reversal of the overall reactions to step 3b in Figure 3 leads to the formation of the corresponding acyl-TDPs, acetyl-TDP and succinyl-TDP, as dead-end intermediates at the active sites of the E1 components. These acyl-TDPs undergo hydrolysis to acetate and succinate. The requirement for the presence of both NADH and TDP in these acyl CoA hydrolyses verified the involvement of E1, E2, and E3 in catalysis, as required by the mechanism in Figure 3. B. Fluoropyruvate as an Alternative Substrate Leading to Acetyl-TDP Pyruvate dehydrogenase from E. coli, the component E1, accepts fluoropyruvate as a substrate and catalyzes its reaction according to either Eq. (4), in the absence of dihydrolipoamide, or Eq. (5), in its presence (11,12):

Reaction (5) leads to the formation of acetyl dihydrolipoamide, just as Eq. (3c) in Figure 3 leads to the formation of acetyl dihydrolipoyl-E2. A key step in the reaction of fluoropyruvate in Eq. (5) may be regarded as a model for Eq. (3c). Reactions (4) and (5) do not require NAD or any other oxidizing agent, yet they lead to products at the acetate oxidation level. This is because fluoropyruvate is at a two-electron-higher oxidation state than pyruvate, by virtue of the fluoro substituent. Shown in Figure 4 is a reasonable mechanistic framework for Reactions (4) and (5). Fluoropyruvate reacts in the same manner as pyruvate through the decarboxylation step to form the enamine intermediate. The en-

8

Frey

Figure 4 Reaction of fluoropyruvate as a substrate for the E1 component of the pyruvate dihydrogenase complex. Fluoropyruvate is an analog of pyruvate and reacts in its place in the early steps of the mechanism shown in Figure 2 for pyruvate decarboxylase. Following decarboxylation, flouride is eliminated from the enamine intermediate, and this leads to enolacetyl-TDP. Upon ketonization, acetyl-TDP is generated at the active site by a process that does not involve a redox step. The acetyl-TDP suffers hydrolysis to acetate and TDP in the absence of an alternative acceptor for the acetyl group. However, dihydrolipoamide captures the acetyl group to form acetyl dihydrolipoamide. The latter process is a model for Reaction (3c) in the overall mechanism shown in Figure 3.

amine is naturally constituted to eliminate fluoride ion, and it does (11) to form enolacetyl-TDP. Ketonization leads directly to acetyl-TDP at the active site of E1, and hydrolysis produces acetate. However, when dihydrolipoamide is present at saturation, hydrolysis does not occur, and acetyl dihydrolipoamide is formed by a process analogous to Reaction (3c) in Figure 3. This last step in the reaction of fluoropyruvate may be regarded as a model for the enzymatic formation of acetyl dihydrolipoyl-E2 in Figure 3. C. Synthesis of Acetyl-TDP The most direct proof of the participation of a putative intermediate in a biochemical or chemical reaction is a demonstration of its formation and transformation into product in the course of the reaction. One step in this process can be accomplished by quenching the reaction in midcourse and proving the presence of the putative intermediate. Experiments such as this


9

are facilitated when the authentic intermediate is available for use in identifying a species isolated from the reaction. In searching for acetyl-TDP as an intermediate in the action of the pyruvate dehydrogenase complex, a sample of acetyl-TDP was required for comparison and identification. The synthesis of acetyl-TDP proved to be more straightforward than originally anticipated; however, its chemical and spectroscopic characterization revealed unexpected and unique properties that eventually strengthened the identification of the same compound isolated in experiments with the pyruvate dehydrogenase complex. Hydroxyethyl-TDP is synthesized by reaction of TDP with acetaldehyde (13). This compound is the logical starting compound for producing acetyl-TDP by an oxidative process. The question of which oxidizing agent is most suitable for hydroxyethyl-TDP revolves around the chemical nature of the group being oxidized and the anticipated chemical properties of the desired product, acetyl-TDP. The hydroxyethyl group in hydroxyethyl-TDP is the only alcoholic group in the molecule, so an oxidizing agent selective for alcohols would be the reagent of choice. Like the 2-acetyl-3,4-dimethylthiazolium ion, acetyl-TDP could be expected to be labile to hydroxide-catalyzed hydrolysis but would be stable in acidic solutions. Therefore, the reagent chosen for the oxidation of hydroxyethyl-TDP was chromic acid. Chromic acid is selective for alcohols, and its mechanism of oxidation leads to dehydrogenation and not oxygenation products, so side reactions such as N-oxidation of the pyrimidine ring are avoided. When used in acidic solutions, the acetyl-TDP produced is stable enough to be purified and characterized (14). Synthetic acetyl-TDP exists in aqueous solutions as a mixture of the three forms shown in Figure 5, a hydrate, the dehydrated acetyl form, and an internal adduct with the 4-amino group of the pyrimidine ring. This mixture was characterized by its 1H NMR spectrum, which showed three sets of resonances for the three forms (14). The equilibrium between the internally cyclized form and the dehyro form was pH dependent because of the ionization of the pyrimidine ring, and this pH dependence introduced a transition step in the pH rate profile for hydrolysis. The unique pH rate profile greatly strengthened the eventual characterization of acetyl-TDP formed in the biological experiments. D. Identification of Acetyl-TDP as an Enzymatic Intermediate Authentic acetyl-TDP potentiated its identification as an intermediate in the action of the pyruvate dehydrogenase complex from E. coli. Reactions of this complex with [2-14C]pyruvate quenched with acid in the steady state yielded a small percentage yield of a radioactive compound that cochromatographed

10

Frey

Figure 5 Structures of the three interconvertible forms of acetyl-TDP in aqueous solutions.

with authentic acetyl-TDP (15). The appearance of this compound depended on the presence of TDP in the reaction, and it was not observed when [1-14C]pyruvate was substituted for [2-14C]pyruvate. Moreover, the pH rate profile for the hydrolysis of the radioactive intermediate proved to be indistinguishable from the unique profile for the hydrolysis of acetyl-TDP (15). The [14C]acetyl-TDP appeared at maximal concentrations in the steady state of the enzymatic reaction and then declined to background at the conclusion of the reaction, as required for an intermediate. Furthermore, the presence of the allosteric inhibitor GTP led to a marked increase in the steady-state concentration of [14C]acetyl-TDP, suggesting that the allosteric inhibitor decreased the rate of acetyl group transfer between E1 and E2. TDP-dependent reactions other than that of the pyruvate dehydrogenase complex have not been examined for the possible participation of acetyl-


11

TDP as a catalytic intermediate. It seems likely that some or al of the various reactions catalyzed by the pyruvate oxidoreductases involve acetyl-TDP, and experiments to test this hypothesis are likely to be fruitful. Moreover, the mechanism of action of phosphoketolase has never been thoroughly studied, and this reaction is also likely to involve the compulsory formation of acetylTDP as an intermediate. IV. A ROLE FOR TDP IN ONE-ELECTRON PROCESSES AND FREE-RADICAL REACTIONS All TDP-dependent enzymes can be assayed by the ferricyanide reduction method because of the universal formation of eneamine intermediates, which are oxidized by ferricyanide. For example, the assay of pyruvate decarboxylase or pyruvate dehydrogenase by this method can be described by Eq. (6) Pyruvate þ H2 O þ FeCN6 3 ! Acetate þ CO2 þ 2FeCN6 4 þ 2Hþ

ð6Þ

These assays must all proceed through free-radical mechanisms because ferricyanide is a compulsory one-electron acceptor. Thus, in reactions that involve the formation of hydroxyethylidene-TDP, the ferricyanide assay must proceed by its oxidation in two one-electron steps by two moles of ferricyanide to acetyl-TDP, which undergoes hydrolysis to TDP and acetate. The implicit intermediate is the one-electron oxidized-radical form of hydroxyethylidene-TDP shown below (16). This intermediate can exist at two protonation levels.

Further one-electron oxidation leads to acetyl-TDP. The formation of this radical in many TDP-dependent reactions is not a biological process and only occurs in the ferricyanide assay. However, in the reactions of pyruvate oxidoreductases it may be a universal intermediate because these enzymes typically carry out one-electron oxidations mediated by iron-sulfur centers. In the case of the pyruvate oxidoreductase from Halobacterium halobium the radical has been observed by EPR and even x-ray crystallography (17–19). The radical is observed as a stable species when pyruvate is added to the enzyme with TDP in the absence of CoA, the cosub-

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Frey

strate. The radical reacts to form acetyl CoA upon addition of CoA. The many resonance forms of this radical may explain its thermodynamic stability, and a detailed spectroscopic analysis by EPR will lead to the identification of the most important resonance forms and the distribution of spin within the molecule (20). The characterization of this radical and related species in other TDP-dependent oxidoreducatases will open a new page in thiamine diphosphate biochemistry.

ACKNOWLEDGMENTS The author gratefully acknowledges the collaboration of graduate students and postdoctoral fellows in his research on thiamine disphosphate mechanisms. These include M. Maldonado, D. C. Speckhard, B. H. Ikeda, M. Apfel, C. A. CaJacob, H. Yang, K. J. Gruys, C. J. Halkides, D. S. Flournoy, C. A. Steginsky, S. Yang, G. R. Gavino, L. S. Leung, and A. Datta.

REFERENCES 1.

R Breslow. Rapid deuterium exchange in thiazolium salts. J Am. Chem. Soc. 79:1762–1763, 1957. 2. TC Bruice, SJ Benkovic. Bioorganic Mechanisms, Vol. II. Benjamin: New York, 1966, pp. 204–214. 3. GL Carlson, GM Brown. The natural occurrence, enzymatic formation, and biochemical significance of a hydroxyethyl derivative of thiamine pyrophosphate. J Biol. Chem. 236:2099–2108, 1961. 4. LO Krampitz, G Gruell, CS Miller, KB Bicking, HR Skeggs, JM Sprague. An active acetaldehyde thiamine intermediate. J Am. Chem. Soc. 80:5893–5894, 1958. 5. H Holzer, K Beaucamp. Nachweis und characterisierung von zwischen Producten der decarboxylierung und oxidation von Pyruvat. Angew. Chem. 71:776–1776, 1958. 6. K Daigo, LJ Reed. Synthesis and properties of 2-acetyl-3,4-dimethylthiazolium iodide. J Am. Chem. Soc. 84:659–662, 1962. 7. GE Lienhard. Kinetics and mechanism of the hydrolysis of 2-acetyl-3,4-dimethylthiazolium ion. J Am. Chem. Soc. 88:5642–5649, 1966. 8. LJ Reed, DJ Cox. Multienzyme complexes. The Enzymes, 3rd ed. 1:213–240, 1970. 9. CA CaJacob, GR Gavino, PA Frey. Pyruvate dehydrogenase complex of Escherichia coli. Thiamine pyrophosphate and NADH dependent hydrolysis of acetyl CoA. J Biol. Chem. 260:14610–14615, 1985. 10. CA Steginsky, PA Frey. Escherichia coli a-ketoglutarate dehydrogenase complex. Thiamine pyrophosphate dependent hydrolysis of succinyl CoA. J. Biol. Chem. 259:4023–4026, 1984.


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11. LS Leung, PA Frey. Fluoropyruvate: an unusual substrate for Escherichia coli pyruvate dehydrogenase. Biochem. Biophys. Res. Communs. 81:274–279, 1978. 12. DS Fluornoy, PA Frey. Pyruvate dehydrogenase and 3-fluoropyruvate: chemical consequence of 2-acetylthiamine pyrophosphate as an acetyl group donor to dihydrolipoamide. Biochemistry 25:6036–6043, 1986. 13. R Kluger, V Stergiopolis, G Gish, K Karimian. Chiral intermediates in thiamin catalysis: Resolution and pyrophosphorylation of hydroxyethylthiamin. Bioorg. Chem. 13:227–234, 1985. 14. KJ Gruys, CJ Halkides, PA Frey. Synthesis and properties of 2-aceylthiamin pyrophosphate: an enzymatic reaction intermediate. Biochemistry 26: 7575– 7585, 1987. 15. KJ Gruys, A Datta, PA Frey. 2-Acetylthiamin pyrophosphate (Acetyl-TPP): pH-rate profile for hydrolysis of acetyl-TPP and isolation of acetyl-TPP as a transient species in pyruvate dehydrogenase catalyzed reactions. Biochemistry 28:9071–9080, 1989. 16. G Barletta, AC Chung, CB Rios, F Jordan. Electrochemical oxidation of enamines related to the key intermediate on thiamin diphosphate dependent enzymatic pathways: evidence for one-electron oxidation via a thiazolium cation radical. J. Am. Chem. Soc. 112:8144–8149, 1990. 17. R Cammack, L Kerscher, D Oesterhelt. A stable free-radical intermediate in the reaction of 2-oxoacid:ferredoxin oxidoreductases of Halobacterium halobium. FEBS Lett. 118:271–273, 1980. 18. S Menon, SW Ragsdale. Mechanism of the Clostridium thermoaceticum pyruvate: ferredoxin oxidoreductase: evidence for the common catalytic intermediacy of the hydroxyethylthiamine pyropyrosphate radical. Biochemistry (July15) 36(28):8484–8494, 1997. 19. E Chabriere, X Vernede, B Guigliarelli, MH Charon, EC Hatchikian, JC Fontecilla-Camps. Crystal structure of the free-radical intermediate of pyruvate: ferredoxin oxidoreductase. Science 294:2559–2563, 2001. 20. PA Frey. Coenzymes and radicals. Science 294:2489–2490, 2001.

2 Mechanistic and Structural Studies on Thiamine Biosynthetic Enzymes Tadhg P. Begley and Steven E. Ealick Cornell University, Ithaca, New York, U.S.A.

I.

INTRODUCTION

The thiamin biosynthetic pathway in B. subtilis is outlined in Figure 1. The thiazole phosphate 4 is formed from glycine 1, cysteine 2, and deoxy-Dxylulose-5-phosphate 3. The pyrimidine phosphate 7 is formed from aminoimidazole ribonucleotide 6. This is then phosphorylated and coupled with the thiazole phosphate 4 to give thiamin phosphate 9. Thiamin pyrophosphate 10, the biologically active form of the cofactor, is formed by a final phosphorylation. In this short review, we will focus on structural and mechanistic studies, rather than a complete review of thiamine biosynthesis, a topic that has recently been reviewed (1).

II.

THIAZOLE PHOSPHATE FORMATION

Thiazole biosynthesis, which requires ThiF, ThiG, ThiS, ThiO, and NifS, has now been reconstituted in a cell-free system, and the function of each of these proteins has been determined (Fig. 2). ThiF catalyzes the adenylation of ThiS 12 to give 13 (2). NifS accepts the sulfur from cysteine to give an active-site persulfide 16 (3), which then adds to 13 (4). Reduction of 14 gives ThiS thio15

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Begley and Ealick

Figure 1

Bacterial thiamine biosynthetic pathway.

carboxylate 15,which functions as an advanced sulfur donor for thiazole biosynthesis (5). ThiO catalyzes the oxidation of glycine (6–8). In a greatly simplified reconstitution of the thiazole biosynthesis, ThiO and glycine can be replaced with glyoxylate and ammonia and ThiF, ThiS, NifS, and cysteine can be replaced by sulfide. In this system, ThiG is the only enzyme required (9). The identification of these partial reactions suggests the mechanism for thiazole formation outlined in Figure 3. In this proposal, glycine is oxidized by ThiO and the resulting glycine imine 17 reacts with the thiocarboxylate of ThiS 15 to give 19. Imine formation followed by tautomerization to 21, thioester hydrolysis to 22, cyclization to 23, double elimination of water to

Mechanisms and Structures of Thiamine Biosynthesis

17

Figure 2 Identification of partial reactions catalyzed by ThiF, ThiS, ThiG, ThiO, and NifS.

Figure 3 Mechanistic proposal for the formation of the thiazole phosphate 4.

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25, and decarboxylation would complete the formation of thiazole phosphate 4. The structures of ThiS and ThiO have been determined (10,11). ThiS has a similar fold to ubiquitin, suggesting that ThiF/ThiS may have been one of the bacterial ancestors to ubiquitin (Fig. 4). X-ray crystallographic analysis revealed that ThiO is a tetrameric protein with identical monomers (10). The ThiO monomer is structurally homologous to sarcosine oxidase (12) and D-amino acid oxidase (13), two other FAD-dependent amine oxidases. ThiO consists of an FAD-binding domain similar to the glutathione reductase 2 family (14) and a substrate binding domain (Fig. 5a). The FAD-binding domain is made up of a mostly parallel six-stranded h-sheet. The sheet is flanked by a six a-helix bundle on one side and by a three-stranded h-sheet and one additional helix on the other side. The substrate binding domain consists of a mixed, eight-stranded h-sheet flanked by three a-helices. The two domains are intertwined with a total of four crossover connections. FAD binds to ThiO in an extended conformation. The FAD forms 14 hydrogen bonds with main chain atoms, three hydrogen bonds with sidechain atoms, and nine additional hydrogen bonds with water molecules. Glycine binds on the re side of the isoalloxazine ring (Fig. 5b). The carboxylate oxygen atoms form two hydrogen bonds with highly conserved Arg302. Residues Glu55 and Arg329 appear to form a lid that closes over the active site after the substrate binds. In addition to the de novo biosynthesis, thiazole alcohol 5 can be salvaged from the growth medium and phosphorylated by thiazole kinase (15).

Figure 4

Comparison of the ThiS and the ubiquitin folds.


19

Figure 5 X-ray structure of ThiO. (a) Structure of the ThiO tetramer. (b) Structure of the ThiO active site showing the isoalloxazine ring of FAD and the substrate glycine.

X-ray crystallographic analysis revealed that thiazole kinase is a trimer of identical subunits (Fig. 6a) (16). The monomer has an a/h structure with homology to the ribokinase family (17). Each monomer contains a central nine-stranded, mostly parallel h-sheet. Five helices flank one side of the sheet and are approximately antiparallel to the strands of the h-sheet. Seven helices are at the opposite side and form a bundle in which three helices abut the central h-sheet. The hydroxyethylthiazole-binding site is located at the interface between two subunits. The ATP binding site stretches out along the C-

Figure 6 X-ray structure of thiazole kinase. (a) Thiazole kinase trimer. (b) Thiazole kinase active site showing the products ADP and thiazole phosphate.

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terminal edge of the h-sheet, placing the terminal phosphate near the substrate binding site. The substrate-binding site is formed by Asn25, Val27, Gly67, Thr68, and Thr194 from one monomer and residues Ala33, Leu37 Pro43, Val 44, and Met45 from an adjacent monomer (Fig. 6b). The side chain of Cys198 is positioned at the bottom of the active site near to the hydroxyl group of the substrate. The ATP binding site is located at the C-terminal ends of h5, h6, h7, and h8. The adenine base is sandwiched between two loops. Asp72 forms hydrogen bonds to the ribose hydroxyl groups, and Asp94, Arg121, Glu126, and Thr168 are involved in ATP phosphate binding. III. PYRIMIDINE PYROPHOSPHATE FORMATION The biosynthesis of the pyrimidine moiety of thiamine has also been reconstituted in a cell free system containing overexpressed ThiC (18). While the reconstitution yield was low, it was possible to determine that the formation of 7 requires S-adenosylmethionine and an additional protein present in the cell free extract (18). A mechanistic proposal for this remarkable rearrangement is outlined in Figure 7. In this proposal, cleavage of the Nglycosidic bond of 6 would give 26 and 27. Ring opening of the ribose 28 followed by a tautomerization and a retro aldol reaction would give 31 and 32. Addition of 32 to the aminoimidazole 27 followed by loss of water, formate, and formaldehyde would give 35. Addition of 31 to aminoimidazole 35 followed by loss of water, ring expansion, and tautomerization would complete the formation of the pyrimidine phosphate 7. HMP-P kinase catalyzes the phosphorylation of pyrimidine phosphate 7. This kinase also catalyzes the phosphorylation of the pyrimidine alcohol 9 in a salvage reaction (19). The X-ray structure of this enzyme has been determined and provides an explanation for this unique dual kinase activity (Fig. 8) (20). X-ray crystallographic analysis revealed that HMP-P kinase is a dimer of identical subunits (Fig. 8a) (20). The monomer has an a/h structure with homology to hydroxyethylthiazole kinase (16) and the ribokinase family (17). Each monomer contains a central eight-stranded, mostly parallel h-sheet. Five a-helices flank one side of the sheet, and the remaining three a-helices are on the opposite side. The structure contains two additional h-strands that form one side of the substrate-binding site. Each monomer has a self-contained active site (Fig. 8b), and the two active sites within the dimer are about 25 A˚ apart. The substrate-binding site is formed by the side chains of Ala18, Val42, Glu 44, Met80, and Val107. A wellordered water molecule is positioned by hydrogen bonds to backbone atoms of Gly11 and Met80 and hydrogen bonds to the substrate. A second active-


21

Figure 7 Mechanistic proposal for the formation of the pyrimidine phosphate 7.

site water molecule is positioned near the first water molecule and the side chains of Asp23 and Cys213. Based on modeling studies, the ATP-binding site is near highly conserved residues Asp187, Lys176, Thr211, and Lys237. Other nearby residues that are highly conserved throughout the ribokinase family include Asp105, Asn139, Glu142, Thr191, and Gly212. Interestingly, the monomers of thiazole kinase and HMP-P kinase are structurally homologous, even though the quaternary structures are entirely different (20). Thiazole kinase is a salvage enzyme and HMP-P kinase is an essential biosynthetic enzyme. In addition, HMP-P kinase catalyzes the phosphorylation of HMP in the salvage pathway. The ATP-binding sites of the two enzymes are structurally similar, while the substrate binding sites are unrelated. The structural homology, limited sequence identity, and similar reactions suggest that these two enzymes may have a common ancestor.

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Figure 8 X-ray structure of HMP-P kinase. (a) Structure of HMP-P kinase dimer. (b) Structure of HMP-P kinase active site with bound ATP and hydroxymethylpyrimidine. An alternate conformation of ATP (not shown) allows for the phosphorylation of HMP-P to HMP-PP.

IV.

THIAMINE PYROPHOSPHATE FORMATION

Thiamine phosphate synthase (ThiE) catalyzes the formation of thiamine monophosphate 9 (21). This reaction proceeds via a dissociative mechanism, and the structure of the enzyme with thiazole phosphate 4, pyrophosphate, and the pyrimidine carbocation has been reported (Fig. 9b) (22,23). The final step in the biosynthetic pathway is catalyzed by thiamine phosphate kinase (ThiL) (24). In addition to the de novo biosynthesis of thiamine phosphate, thiamine alcohol can be salvaged from the growth medium and converted to thiamine phosphate. This reaction is catalyzed by thiamine kinase (YcfN in E.coli) (25).

Figure 9 X-ray structure of thiamine phosphate synthase. (a) Tertiary structure of thiamine phosphate synthase. (b) Active site of thiamine phosphate synthase showing the pyrimidine carbocation intermediate, thiazole, and pyrophosphate.


23

X-ray crystallographic analysis revealed that thiamine phosphate synthase is an a/h protein with a (ha)8 barrel fold (Fig. 9a) (26). An additonal Nterminal a-helix caps the bottom of the barrel, and an additional short helix is located in the loop after h8. The active site is located at the top of the barrel near the C-terminal ends of the h-strands. Seven of the eight ha loops, 2–8, form the active site. Loops 2–5 are involved primarily with HMPPP interactions, and loops 6–8 are involved primarily in thiazole phosphate interactions. About 13 highly conserved residues form the active site (Fig. 9b). Arg59, Lys61, Asn9, Lys 159, and Mg+2 form interactions with the h-phosphate group of HMP-PP, while the a-phosphate group interacts with Ser130A, Lys159, Mg+2, and four water molecules. The PPi Mg+2 is ligated to Asp93, Asp112, and two water molecules. The pyrimidine forms two hydrogen bonds with Gln57. The phosphate group of thiazole phosphate forms hydrogen bonds with Thr156, Thr158, Gly188, Ile208, and Ser 209. The thiazole ring forms several van der Waals contacts. The structure of the S130A mutant treated with substrates revealed unexpectedly separate thiazole, pyrimidine, and pyrophosphate moieties, thus providing structural evidence for a carbocation intermediate (22). V. ENZYMATIC SYNTHESIS OF THIAMINE PYROPHOSPHATE The availability of overexpression strains for all of the later enzymes involved in thiamine pyrophosphate biosynthesis and salvage makes possible a facile synthesis from the readily synthesized pyrimidine 39 and thiazole alcohol 5 (27). This may be of use for the preparation of isotopically labeled thiamine pyrophosphate, a useful NMR and ESR probe for thiamine-utilizing enzymes. VI.

MECHANISM OF BACIMETHRIN TOXICITY

Using the later enzymes on the thiamine biosynthetic pathway, it was possible to demonstrate that bacimethrin 40, an antibiotic whose activity can be re-

Figure 10 An efficient enzymatic synthesis of thiamine pyrophosphate 10.

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Figure 11 The antibiotic bacimethrin is converted to methoxythiamine pyrophosphate by the thiamine pyrophosphate biosynthetic enzymes.

versed by thiamine, is converted to methoxythiamine pyrophosphate 41. This reaction occurs six times faster than the rate of conversion of 39 to thiamine pyrophosphate (10, Figs. 10 and 11). This thiamine analog is an antivitamin and inhibits transketolase, deoxy-D-xylulose-5-phosphate synthase, and aketoglutarate dehydrogenase, three of the seven thiamine-requiring enzymes in E. coli (28). VII.

THIAMINE DEGRADATION

Thiaminase I catalyzes the replacement of the thiazole moiety of thiamine by a variety of nucleophiles (Fig. 1). The structure of the enzyme with a bound mechanism-based inactivating agent is shown in Figure 12. This structure supports the mechanism outlined in Figure 13 (29,30). In this mechanism,

Figure 12 X-ray structure of thiaminase I. (a) Tertiary structure of thiaminase I. (b) Active site of thiaminase I with a bound mechanism-based inactivating agent.


Figure 13

25

Mechanistic proposal for thiaminase I.

Cys113 adds to C6 of the pyrimidine to give 40. Loss of the thiazole from the resulting anion gives 41. Reversal of these two steps, using the added nucleophile, completes the substitution reaction. The biological function of thiaminase I has not yet been determined. X-ray crystallographic analysis revealed that thiaminase I contains two intertwined a/h domains (Fig. 12a) (29). One domain is made up primarily from the N-terminal half of the chain and the other from the C-terminal half. Each domain consists of a three-layer aha sandwich, and the two domains are structurally similar to each other. The overall structure is homologous to that of the periplasmic-binding proteins (31), which provide specific binding sites for various small molecules, and the transferrins (32). The active site of thiaminase I is located in a cleft between the two domains and is in a location similar to the small molecule–binding site of the periplasmic-binding proteins. The thiamine phosphate–binding site is shown in Figure 12b. The binding site was confirmed by determining the structure of thiaminase I in complex with the mechanism-based inhibitor 4-amino-6-chloro-2-methylpyrimidine. The active site cleft is formed by residues 16–20, 48–50 and 64–66 from the N-terminal domain and residues 158–164, 213–224, and 238–241 from the C-terminal domain. Six tyrosine residues, Tyr16, Tyr18, Tyr50,

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Tyr222, Tyr239, and Tyr270, form a collar of residues within the active-site cleft. Cys113 is located at the bottom of the cleft and is the site of covalent attachment of the inhibitor. Glu241 is located nearby and is believed to serve as an active site base to increase the nucleophilicity of Cys113. VIII.

SUMMARY

The biosynthesis of thiamine pyrophosphate in bacteria is a complex process requiring at least 11 biosynthetic and salvage enzymes. The later steps in the pathway are now well understood: considerable progress has been made with the thiazole forming reactions, and the successful reconstitution of the pyrimidine biosynthesis is a promising first step toward elucidating the mechanism of this remarkable reaction. Six of the proteins involved in thiamine pyrophosphate biosynthesis and degradation have been structurally characterized. REFERENCES 1.

2.

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8.

TP Begley, D Downs, S Ealick, F McLafferty, DV Loon, S Taylor, N Chiu, J Chiu, C Kinsland, J Reddick, J Xi. Thtiamin biosynthesis in prokaryotes. Arch Microbiol 171:293–300, 1999. SV Taylor, NL Kelleher, C Kinsland, H-J Chiu, CA Costello, AD Backstrom, FW McLafferty, TP Begley. Thiamin biosynthesis in Escherichia coli. Identification of the ThiS thiocarboxylate as the immediate sulfur donor in the thiazole formation. J Biol Chem 273:16555–16560, 1998. CT Lauhon, R Kambampati. The iscS gene in Escherichia coli is required for the biosynthesis of 4-thiouridine, thiamin, and NAD. J Biol Chem 275:20096– 20103, 2000. J Xi, Y Ge, C Kinsland, FW McLafferty, TP Begley. Biosynthesis of the thiazole moiety of thiamin in Escherichia coli: Identification of an acyldisulfidelinked protein–protein conjugate that is functionally analogous to the ubiquitin/E1 complex. Proc Natl Acad Sci USA 98:8513–8518, 2001. TP Begley, J Xi, C Kinsland, S Taylor, F McLafferty. The enzymology of sulfur activation during thiamin and biotin biosynthesis. Curr Op Chem Biol 3:623– 629, 1999. V Job, GL Marcone, MS Pilone, L Pollegioni. Glycine oxidase from Bacillus subtilis. Characterization of a new flavoprotein. J Biol Chem 277:6985–6993, 2002. V Job, G Molla, MS Pilone, L Pollegioni. Overexpression of a recombinant wild-type and His-tagged Bacillus subtilis glycine oxidase in Escherichia coli. Eur J Biochem 269:1456–1463, 2002. Y Nishiya, T Imanaka. Purification and characterization of a novel glycine oxidase from Bacillus subtilis. FEBS Lett 438:263–266, 1998.

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J-H Park, P Dorrestein, TP Begley. Reconstitution of the biosynthesis of the thiazole moiety of thiamin. Unpublished, 2002. EC Settembre, P Dorrestein, J-H Park, A Augustine, TP Begley, SE Ealick. Structural and mechanistic studies on ThiO, a glycine oxidase essential for thiamin biosynthesis in Bacillus subtilis. Biochemistry 42(10):2971–2981, 2003. C Wang, J Xi, TP Begley, TP Begley, TP Begley, LK Nicholson. Solution structure of ThiS and implications for the evolutionary roots of ubiquitin. Nat Struct Biol 8:47–51, 2000. P Trickey, MA Wagner, MS Jorns, FS Mathews. Monomeric sarcosine oxidase: structure of a covalently flavinylated amine oxidizing enzyme. Structure 7:331–345, 1999. A Mattevi, MA Vanoni, F Todone, M Rizzi, A Teplyakov, A Coda, M Bolognesi, B Curti. Crystal structure of D-amino acid oxidase: a case of active site mirror-image convergent evolution with flavocytochrome b2. Proc Natl Acad Sci USA 93:7496–7501, 1996. O Dym, D Eisenberg. Sequence–structure analysis of FAD-containing proteins. Protein Sci 10:1712–1728, 2001. Y Zhang, SV Taylor, H-J Chiu, TP Begley. Characterization of the Bacillus subtilis thiC operon involved in thiamine biosynthesis. J Bacteriol 179:3030, 1997. N Campobasso, I Matthews, TP Begley, SE Ealick. Crystal structure of 4methyl-5-h-hydroxyethylthiazole kinase from B. subtilis. Biochemistry 39:7868– 7877, 2000. JA Sigrell, AD Cameron, TA Jones, SL Mowbray. Structure of Escherichia coli ribokinase in complex with ribose and dinucleotide determined to 1.8-A˚ resolution: insights into a new family of kinase structures. Structure 6:183–193, 1998. R Mehl, B Lawhorn, TP Begley. Reconstitution of the biosynthesis of the pyrimidine moiety of thiamin. In preparation, 2002. JJ Reddick, C Kinsland, LA Petersen, ME Winkler, DM Downs, TP Begley. Overexpression, purification and characterization of two kinases involved in the biosynthesis of thiamin. Tetrahedron 54:15983–15991, 1998. G Cheng, EM Bennett, TP Begley, SE Ealick. Crystal structure of 4-amino-5hydroxymethyl-2-methylpyrimidine phosphate kinase from Salmonella typhimurium at 2.3-A˚ resolution. Structure 10:225–235, 2002. A Backstrom, A McMordie, TP Begley. Biosynthesis of thiamin (I): the function of the thiE gene product. J Am Chem Soc 117:2351–2352, 1995. DH Peapus, H-J Chiu, N Campobasso, JJ Reddick, TP Begley, SE Ealick. Structural characterization of the enzyme–substrate, enzyme–intermediate, and enzyme–product complexes of thiamin phosphate synthase. Biochemistry 40: 10103–10114, 2001. JJ Reddick, R Nicewonger, TP Begley. Mechanistic studies on thiamin phosphate synthase: evidence for a dissociative mechanism. Biochemistry 40:10095– 10102, 2001. E Webb, D Downs. Characterization of thiL, encoding thiamin-mono-

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Begley and Ealick phosphate kinase, in Salmonella typhimurium. J Biol Chem 272:15702–15707, 1997. J Melnick, E Lis, H Mori, C Kinsland, J Perkins, G Schyns, A Osterman, TP Begley. Identification of two new thiamin salvage enzymes. Unpublished, 2002. H-J Chiu, J Reddick, TP Begley, SE Ealick. The structure of thiamin phosphate synthase from Bacillus subtilis. Biochemistry 38:6460–6470, 1999. J Melnick, TP Begley. The enzymatic synthesis of thiamin pyrophosphate. In preparation, 2002. JJ Reddick, S Saha, J Lee, JS Melnick, J Perkins, TP Begley. The mechanism of action of bacimethrin, a naturally occurring thiamin antimetabolite. Bioorganic Med Chem Lett 11:2245–2248, 2001. N Campobasso, C Costello, C Kinsland, TP Begley. Crystal structure of thiaminase I from Bacillus thiaminolyticus at 2.0-A˚ resolution. Biochemistry 37:15891–15989, 1998. C Costello, N Kelleher, M Abe, F McLafferty, TP Begley. Mechanistic studies on thiaminase I(2): overexpression and identification of the active-site nucleophile. J Biol Chem 271:3445–3452, 1996. JC Spurlino, GY Lu, FA Quiocho. The 2.3-A˚ resolution structure of the maltose- or maltodextrin-binding protein, a primary receptor of bacterial active transport and chemotaxis. J Biol Chem 266:5202–5219, 1991. S Bailey, RW Evans, RC Garratt, B Gorinsky, S Hasnain, C Horsburgh, H Lindley, PF Lindley, A Mydin, R Sarra, et al. Molecular structure of serum transferrin at 3.3-A˚ resolution. Biochemistry 27:5804–5812, 1988.

3 Studies on the Structure and Function of Thiamine Pyrophosphokinase Jing-Yuan Liu, Robert A. Harris, and Thomas D. Hurley Indiana University School of Medicine, Indianapolis, Indiana, U.S.A. David E. Timm Eli Lilly and Company, Indianapolis, Indiana, U.S.A.

I. INTRODUCTION Thiamine, also known as vitamin B1, is biologically active only when it is converted into thiamine pyrophosphate (TPP) (1) by thiamine pyrophosphokinase (TPK) (2). In humans, TPP serves as a prosthetic group for four metabolically important enzymes: branched chain a-keto acid dehydrogenase, pyruvate dehydrogenase, a-ketoglutarate dehydrogenase, and transketolase. The common chemical feature of these enzymes involves the transfer of an activated aldehyde unit. Because thiamine plays a control role in intermediary metabolism, it is important for the body to maintain a normal steady-state concentration of intracellular TPP. Thiamine deficiency causes brain damage (3) in Wernicke–Korsakoff syndrome (4) and thiamineresponsive megaloblastic anemia (5). Thiamine deficiency is also a frequent complication of alcoholism (6,7). It is reported that TPK activity in supernatants of rat brain tissues is significantly reduced in response to acute or chronic ethanol administration as compared with control rats (8).

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The structure of mouse thiamine pyrophosphokinase (mTPK) (9) as both an apoenzyme and thiamine-bound form has been solved (10). The enzyme is a homodimer. Each monomer contains an N-terminal a/h-domain and a C-terminal h-sandwich domain. Thiamine is bound in the dimer interface and adopts an F-conformation. Little is known about the catalytic mechanism and the regulation of TPK. Further structural and enzymological study will help to elucidate the function and catalytic mechanism of this crucial enzyme.

II. MATERIALS AND METHODS A. Expression and Kinetics The coding region for mouse TPK was cloned into the expression vector pET28a and expressed as a his-tag fusion protein in E. coli, as previously described (10). The recombinant protein was purified by nickel-chelate chromatography (Sigma) and concentrated to 11.2 mg/mL and stored in 150 mM NaCl at 80oC. The N-terminal his tag (20 AA’s) was not removed prior to further characterization. Kinetic activity assays were performed using a coupled enzyme system (11) that included mouse TPK, myokinase (Sigma), pyruvate kinase (Sigma), and lactate dehydrogenase (Sigma). Each mole of AMP generated by mTPK causes the oxidation of 2 moles of NADH. The disappearance of NADH was monitored at 340 nm. The reaction was initiated by adding 56 Ag TPK enzyme to the assay mixture (1 mL total). One unit is defined by the amount of enzyme required to convert 1 nmol thiamine to TPP/min. B. Crystallization and Crystal Soaking Crystals were prepared by vapor diffusion, as described previously (10). Hanging drops were set up using an equal volume of an 11.2 mg/mL TPK solution and well solution, which contains 1.8–2.1 M ammonium sulfate, 0.1 M Hepes and 2–4% PEG400, pH 7.1 at room temperature. Crystals developed to full size within two weeks. TPK crystals were then soaked overnight in a well solution containing 10 mM pyrithiamine and 10 mM ATP for the pyrithiamine-bound structure or well solutions containing 10 mM TPP for TPP-bound structure, respectively. C. Structure Determination and Refinement Diffraction data for mTPK crystals were collected with a Rigaku RU200 rotating-anode X-ray generator utilizing an R-axis IIc image plate detector. The raw intensity data were reduced using HKL (v1.96.0) and merged and

Structure and Function of TPK

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Table 1 Crystallographic Data and Refinement Statistics Data collection Resolution (A˚) Space group Cell constants a = b, c (A˚) a = h, g (j) Measured reflections Unique reflections Completeness (%) Rsym (%) Refinement Rwork/Rfree (%) Number of atoms/asymmetric unit Nonhydrogen protein atoms Solvent molecules Root-mean-square deviation Bond length (A˚) Bond angles (j)

TPP

Pyrithiamine

30.00–1.70 P3121

30.00–1.95 P3121

89.9, 141.4 90, 120 184,997 67,389 92.6 3.4 (44.5)

89.9, 141.2 90, 120 190,670 44,429 91.1 5.0 (30.9)

22.4/23.9

22.1/25.6

3,394 446

4,015 473

0.008 1.4

0.007 1.3

scaled by SCALEPACK (12). The intensities output by HKL were converted into structure factors using the program package CCP4 (13). The initial phase information was acquired from the mTPK apoenzyme structure. Models were manually adjusted and refined by CNS (v1.0) (14). Pyrithiamine coordinates were created by CORINA (15,16). Thiamine pyrophosphate coordinates were acquired from the protein data bank. Statistics for crystallographic data and refinement are shown in Table 1. III. RESULTS AND DISCUSSION A. Kinetic Studies TPK activities were measured at room temperature in four different buffer systems of 0.1 M, pH 7.4 each (Fig. 1). Phosphate buffer gave the lowest TPK activity. Activity measured in Hepes buffer were about six times that obtained in phosphate buffer, and assays measured in Tris-HCl and ACES buffers were about five-fold higher than in phosphate buffer. These observations suggest that inorganic phosphate may have an inhibitory effect on mTPK. It is possible that phosphate inhibits the enzyme by competing for the ATP-binding pocket. This possibility is supported by the fact that sulfate ions, which also have an inhibitory effect on mTPK (data not shown), were

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Figure 1 mTPK activity in four different buffers (0.1 M, pH 7.4): Tris-HCl, phosphate, HEPES, and ACES. Activity was measured with a coupled enzyme system at 23oC. Phosphate buffer has a five- to six-fold lower activity as compared with others. One unit is defined by the amount of enzyme required to convert 1 nmol thiamine to TPP per min.

identified in the presumed ATP-binding pocket. Because sulfate and phosphate have similar structures and similar molecular size, it is possible that both inhibit the enzyme by binding and competing with ATP for its binding pocket. Most previous kinetic studies performed on TPK used phosphate buffers because of its high buffer capacity (8,17) or used buffers containing inorganic phosphate or sulfate ions (18–21). The kinetic parameters derived from these studies generated very high Km (from 59 to 5.8 mM) for ATP. Yet the physiological concentration of ATP, for example, in brain, is about 3 mM (22). This raises the question of how TPK catalyzes the reaction to generate thiamine pyrophosphate from ATP. The current finding, that phosphate and sulfate both inhibit TPK, suggests that the kinetic parameters measured previously of TPK may need to be reassessed. Alcoholics often develop thiamine deficiency and neurodegenerative disease. It was reported that TPK activity was inhibited when ethanol was added to the supernatant of different regions of rat brain tissues (8). However, ethanol did not inhibit the recombinant mouse TPK at a concentration of 25 mM, which is equivalent to 110 mg/dL blood alcohol level (Fig. 2). Activity assays were performed in 0.2 M Tris-phosphate and Tris-HCl buffers, pH7.4, respectively. mTPK activity was not affected by ethanol in either buffer system tested. Furthermore, no significant inhibition was observed up to an ethanol concentration of 100 mM (data not shown).


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Figure 2 Ethanol effect on mTPK activity. Assays with or without 25 mM alcohol were performed in Tris-phosphate buffer and Tris-HCl buffer (both 0.2 M, pH7.4) at 22oC, respectively. No alcohol inhibition on the recombinant enzyme was observed.

Mouse TPK was found to be expressed mainly in liver and kidney when the cDNA was untilized as a probe for Northern blots (9). However, human TPK, which has an 89% sequence identity to mouse TPK, was expressed in a wide variety of tissues, with very low amounts, if any, in brain (23). Considering the importance of TPP for metabolism in brain tissues and the previous reports of TPK activity in brain extracts, the finding that ethanol does not inhibit recombinant mTPK suggests that there may be another isoform of TPK specifically expressed in the brain, which can be inhibited by ethanol. It is reported that both pig TPK from heart and brain are homodimers. However, their molecular weights are different (24–26). This evidence seems to support the isoenzyme idea of TPK. It is also possible that TPP phosphatase can be activated by ethanol and results in an apparent inhibition of TPK in the supernatant of brain tissues.

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In addition to ethanol, TPP, AMP, and many thiamine analogs, such as oxythiamine and pyrithiamine, are known inhibitors of TPK. TPP and AMP are products of TPK catalysis and probably inhibit TPK by occupying the thiamine and ATP-binding pockets. Oxythiamine and pyrithiamine are the two most extensively studied thiamine analogs. Oxythiamine differs from thiamine by replacing the 4V-NH2 group of thiamine with a hydroxyl group. Pyrithiamine replaces the thiazolium ring of thiamine with a pyridinium ring (Fig. 3). That is, pyrithiamine substitutes the sulfur of thiamine thiazolium ring with a CHCH group. To understand the mechanism of inhibition for TPP and pyrithiamine, X-ray crystallography was used to determine structures of TPK complexed with TPP and with pyrithiamine at 1.70-A˚ and 1.95A˚ resolution, respectively. B. Thiamine Pyrophosphate (TPP) Structure The TPP-complexed structure shows that TPK binds TPP in the thiaminebinding pocket (Figs. 4A, 5A). The overall structure for the TPP-bound form of TPK is very similar to the thiamine-bound form. TPP was found at the hydrophobic interface of the two subunits, and residues from both subunits contribute to TPP binding. The similarities between TPP- and thiaminebinding suggest that only minimal changes in structure accompany the conversion of substrates into products.

Figure 3 Substrates, products, and inhibitors of TPK.


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Figure 4 2Fo-Fc maps for TPP and pyrithiamine. (A) Electron density map for thiamine pyrophosphate contoured at 1 standard deviation. (B) Electron density map for pyrithiamine contoured at 1 standard deviation.

One difference between thiamine- and TPP-binding concerns Asp117, where its hydrogen bond to the 4V-amino group in the TPP structure is weaker than that in the thiamine-complexed structure, due to the fact that the carboxyl oxygen is further away from the 4V-amino group (3.5 A˚ as compared to 2.5 A˚ in the thiamine structure). The side chain of Gln154, which is bent away from thiamine in the thiamine-bound structure, extends toward TPP and, together with the peptide nitrogen atom from Asp120 and the side chains of Gln154 and Arg151, hydrogen-bonds to the pyrophosphate group of TPP (Fig. 5A). Comparison of the TPP binding environment and its conformation between TPK and TPP-dependent enzymes provides interesting insight (Figs. 5A, 5B). TPP in both mTPK and TPP-dependent enzymes is in a hydrophobic cleft of two subunits. Aromatic residues are usually recruited in pyrimidine ring binding. In mTPK, this residue is Trp; in branched-chain

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Figure 5 Thiamine pyrophosphate (TPP) binding in mouse thiamine pyrophosphokinase (mTPK) and branched-chain a-keto acid dehydrogenase E1 component (BCKDH). TPP and Mg2+are shown in dark gray. Nitrogen atoms are in black; oxygen atoms in gray and carbon atoms in light gray. Both TPP molecules were found at the interface of A and B subunits and residues from both subunits contribute to TPP binding. Residues from B subunit are marked by a prime symbol. (A) TPP binding in mTPK. W222V interacts with the pyrimidine ring through stacking interactions. The carbonyl oxygens of T237V and Q116 stabilize the positively charged thiazolium ring nitrogen by dipolar interactions. The side chain of D117 hydrogen-bonds to the 4Vamino group of the pyrimidine ring. The amino groups of D120, R151, and Q154 stabilize the pyrophosphate group of TPP by neutralizing its negative charges. (B) TPP binding in BCKDH. Y102V stacking interacts with the TPP pyrimidine ring. The sidechain oxygen of S162 hydrogen bonds with the TPP 4V-NH2 group. The side chain of E76V hydrogen-bonds with the TPP N1V atom. The nitrogen atoms from side chains of Q112, R114, R220, and H291 and from the main chain of G194 (not shown) hydrogenbond the oxygen atoms of the TPP phosphate groups. The figures were made using the Swiss PDB Viewer and POV-Ray for Windows. BCKDH coordinates were accessed from the Protein Data Bank with the code 1DT. (From Ref. 27.)


37

a-keto acid dehydrogenase E1 component (BCKDH) (27), this residue is Tyr. The functional difference of TPP between TPK and TPP-dependent enzymes can be explained through the differences in their structural basis for TPP binding. First, TPP-dependent enzymes bind pyrophosphate group more tightly, due to the fact that more and stronger hydrogen bonds exist (Fig. 5B; see more detail in the figure legend). This is not surprising, because TPP serves as a prosthetic group in TPP-dependent enzyme while it serves as a product feedback inhibitor in TPK. TPK has mild affinity for TPP, so it would be released from the protein under certain conditions and TPK would be able to catalyze a new round of reaction. Second, in mTPK, TPP phosphate groups are in a solvent-accessible space, where the catalytic reaction occurs. However, in TPP-dependent enzymes, the C2 atom of the thiazolium ring is the only atom that is solvent accessible. The C2 atom is the active center of TPP-dependent enzymes, and it is critical for the C2 atom to be accessible by substrates. Last, hydrogen bonds between the conserved Ser and the 4V-NH2 group of TPP and between Glu and the N1V atom of TPP in TPP-dependent enzymes are missing in TPK. These two hydrogen bonds are functionally important in activating the C2 atom and, thus, are conserved among TPP-dependent enzymes (28). The reaction catalyzed by TPK does not involve the deprotonation and activation of the C2 atom. Thus, these two hydrogen bonds are functionally meaningless to TPK. TPK binds TPP in the F-conformation. In contrast, TPP in all TPPdependent enzyme structures is bound in the V-conformation (Fig. 6). The terms F- and V-conformation are a description of the orientation of the two

Figure 6 Superposition of thiamine pyrophosphate from mTPK and from BCKDH, an enzyme that utilizes TPP as a prosthetic group. TPP in mTPK adopts a low-energy F-conformation (in black), while the cofactor TPP adopts a highenergy V-conformation (in gray).

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Figure 7 Numerical assignment of thiamine ring atoms.

planar ring systems of TPP with respect to each other, defined by the parameters AT and AP (29). AT is defined by the dihedral angle between C5V-C (3, 5V) and N3-C2; AP is the dihedral angle between N3-C (3, 5V) and C5V-C4V (Fig. 7). AT and AP are 6.8j and 83.8j in an ideal F-conformation and 103.5j and 63.9j in an ideal V-conformation. The AT and AP values for TPP found in mTPK are 2j and 81.5j, respectively. The V-conformation is a high-energy state but is functionally important for catalysis in TPP-dependent enzymes. C. Pyrithiamine-Complexed Structure The TPK inhibitor pyrithiamine was frequently used to induce thiamine deficiency in rodents due to its ability to deplete thiamine and produce the neurological syndrome of thiamine deficiency (30,31). Both oxythiamine and pyrithiamine can induce thiamine deficiency. However, the symptoms are different. Oxythiamine produces thiamine-deficiency symptoms such as anorexia and lethargy but no neurological symptoms. In contrast, pyrithiamine produces primarily neurological symptoms (32). Oxythiamine does not deplete thiamine, while pyrithiamine does (33). It is clear that oxythiamine is a competitive inhibitor of TPK and can be phosphorylated by TPK (34). Since the thiamine 4V-NH2 group, which is important in deprotonating thiamine pyrophosphate C2 atom and activating TPP-dependent enzymes, is missing in oxythiamine; the phosphorylated form of oxythiamine can bind to TPP-dependent enzymes but yields an inactive enzyme (35). Pyrithiamine is a more potent inhibitor of TPK than oxythiamine, but its inhibition mechanism is unclear. To understand how pyrithiamine interacts with TPK and its mechanism of inhibition, X-ray diffraction data were collected using crystals soaked with pyrithiamine and ATP. Pyrithiamine was identified in the active site (Table 1). The overall structure of pyrithiamine-bound mTPK is similar to the thiamine-complexed structure, with an rmsd of main-chain and sidechain atoms of 0.52 A˚. Pyrithiamine is not phosphorylated, and it adopts the


39

low-energy F-conformation, similar to thiamine (Fig. 8). Furthermore, the hydroxylethyl group of pyrithiamine is relatively flexible and is open to a solvent-accessible space of the cleft. The residues that contribute to pyrithiamine binding are similar to those that contact thiamine and the rmsd of the main-chain and side-chain atoms of these residues in the two structures is 0.08 A˚. Residues 116–119 interact with the pyridinium ring in this structure, while they contribute to thiazolium ring interactions in the thiamine-complexed structure. The carbonyl oxygen of Gln116 is closer to the N1 atom of pyrithiamine, and thus the dipolar interaction between the two atoms is stronger as compared with that of the thiamine-complexed structure (3.05 A˚ versus 3.50 A˚). The absence of the bulky and electron-rich sulfur in pyrithiamine is the possible reason that made the close distance between the carbonyl oxygen of Gln116 and N1 atom stereochemically and electrochemically allowed. Another difference involves residue Asp91. The side chain of Asp91 bends toward the hydroxyl oxygen of thiamine and forms a hydrogen bond in thiamine-bound structure. However, in pyrithiamine-complexed structure, the side chain of Asp91 extends away from the hydroxyl oxygen of the pyrithiamine, and no hydrogen bond formed.

Figure 8 Pyrithiamine binding in mTPK. Pyrithiamine is found in the thiaminebinding pocket. Thiamine (gray) is superimposed on pyrithiamine (black). Both of them adopt an F-conformation. Asp91 of the thiamine-bound structure (gray) is superimposed on that of the pyrithiamine-bound structure, to show the difference in this residue between the two structures. The figures were made by using the Swiss PDB Viewer and POV-Ray for Windows.

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ATP could not be located bound to mTPK. Instead, two sulfates were found in what we believe is the ATP-binding pocket. Currently, it is still controversial whether pyrithiamine can be phosphorylated by TPK (31). The pyrithiamine in the structure was not phosphorylated. However, the structural data do not provide enough evidence for why pyrithiamine could not be phosphorylated. The lack of pyrithiamine phosphorylation in this structure may be due to the inability of ATP to bind to this crystal form. Thus, more studies on the TPK catalytic mechanism are needed to determine whether pyrithiamine can be phosphorylated by TPK. Attempts to obtain crystals with MgATP bound to TPK have been unsuccessful. Soaking with 0.1 M MgATP could not establish the binding, whereas soaking with 0.2 M MgATP dissolved the crystals. Soaking another crystal form grown in 21% ethanol, 0.1 M glycine at pH 10.0 had similar results. It is possible that TPK catalyzes pyrophosphate transfer from ATP to thiamine by an ordered mechanism and that MgATP binds to TPK first and thiamin second (21). However, the crystallographic data challenge this notion. Screening crystals of mTPK with a high concentration of thiamine, AMP-CPP and MgCl2 or TPP, MgCl2, and AMP have been tried, and new crystal forms have appeared in some of the conditions. Further studies on the structural and kinetic properties of catalysis in TPK are warranted and currently under investigation.

ACKNOWLEDGMENTS The authors thank Paul Blair for useful discussion and help with the TPK kinetic assay, John Hawes for the help with TPK cloning, and Paresh Sanghani and Sam Perez-Miller for their help in graphic preparation. The work was supported by NIH grants DK19259 (RAH) and DK 54738 (DET) and a pilot grant from the Indiana Alcohol Research Center (DET and TDH). REFERENCES 1.

2.

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H Mitsuda, Y Takii, K Iwami, K Yasumoto. Purification and properties of thiamine pyrophosphokinase from parsely leaf. J Nutr Sci Vitaminol (Tokyo) 21:103–115, 1975. IM Artsukevich, AI Voskoboev, M Iu Ostrovskii. [Purification and several properties of thiamine pyrophosphokinase from rat liver]. Vopr Med Khim 23:203–210, 1977. C Harper, J Kril. An introduction to alcohol-induced brain damage and its causes. Alcohol Alcohol Suppl 2:237–243, 1994.

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CK Singleton, PR Martin. Molecular mechanisms of thiamine utilization. Curr Mol Med 1:197–207, 2001. G Rindi, C Patrini, U Laforenza, H Mandel, M Berant, MB Viana, V Poggi, AN Zarra. Further studies on erythrocyte thiamin transport and phosphorylation in seven patients with thiamin-responsive megaloblastic anaemia. J Inherit Metab Dis 17:667–677, 1994. D Hell, P Six. [Thiamine, riboflavine and pyridoxine supply in chronic alcoholism (author’s transl)]. Dtsch Med Wochenschr 102:962–966, 1997. CM Leevy. Thiamin deficiency and alcoholism. Ann NY Acad Sci 378:316–326, 1982. G Rindi, L Imarisio, C Patrini. Effects of acute and chronic ethanol administration on regional thiamin pyrophosphokinase activity of the rat brain. Biochem Pharmacol 35:3903–3908, 1986. K Nosaka, M Onozuka, H Nishino, H Nishimura, Y Kawasaki, H Ueyama. Molecular cloning and expression of a mouse thiamin pyrophosphokinase cDNA. J Biol Chem 274:34129–34133, 1999. DE Timm, J Liu, LJ Baker, RA Harris. Crystal structure of thiamin pyrophosphokinase. J Mol Biol 310:195–204, 2001. HU Bergmeyer. Methods of Enzymatic Analysis. New York: Academic Press, pp 2127–2131, 1974. Z Otwinowski, W Minor. Processing of X-ray Diffraction Data Collected in Oscillation Mode. In: J.C.W. Carter, R.M. and Sweet, eds. Methods in Enzymology. Vol. 276. New York: Academic Press, 1997, 307–326. Collaborative Computing Project No. 4 The CCP4 suite: programs for protein crystallography. Acta Crystallographica D50:760–763, 1994. AT Brunger. X-ray crystallography and NMR reveal complementary views of structure and dynamics. Nat Struct Biol 4(suppl):862–865, 1997. J Gasteiger, C Rudolph, J Sadowski. Automatic generation of 3D-atomic coordinates for organic molecules. Tetrahedron Comp Method 3:537–547, 1990. J Sadowski, C Rudolph, J Gasteiger. The Generation of 3D models of host–guest complexes. Analytica Chimica Acta 265:233–241, 1992. C Patrini, G Rindi. An improved method for the electrophoretic separation and fluorometric determination of thiamine and its phosphates in animal tissues. Int J Vitam Nutr Res 50:10–18, 1980. JW Peterson, CJ Gubler, SA Kuby. Partial purification and properties of thiamine pyrophosphokinase from pig brain. Biochim Biophys Acta 397:377– 394, 1975. SM Sanioto, H Reinauer, S Hollmann. Thiamine pyrophosphokinase activity in liver, heart and brain crude extracts of control and thiamine deficient rats. Int J Vitam Nutr Res 47:315–324, 1977. AI Voskoboyev, YM Ostrovsky. Thiamin pyrophosphokinase: structure, properties, and role in thiamin metabolism. Ann NY Acad Sci 378:161–176, 1982. Clark J. Gubler, G Fleming, SA Kuby. Thiamin pyrophosphokinase (ATP thiamin pyrophosphotransferase) E.C. 2.7.6.2: Purification, properties and function. In: H. Bisswanger, and A. Schellenberger, eds. Biochemistry and Phys-

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Liu et al. iology of Thiamin Diphosphate Enzyme. Prienam Chiemsee, Germary: Intemann, 557–569, 1996. AG Nasello, R Depiante, M Tannhauser, I Izquierdo. Effect of morphine on the RNA and ATP concentration of brain structures of the rat. Pharmacology 10:56–59, 1973. K Nosaka, M Onozuka, N Kakazu, S Hibi, H Nishimura, H Nishino, T Abe. Isolation and characterization of a human thiamine pyrophosphokinase cDNA. Biochim Biophys Acta 1517:293–297, 2001. M Hamada. [Thiamine pyrophosphokinase of pig heart muscles]. Seikagaku 41:837–849, 1969. Y Wakabayashi, A Iwashima, Y Nose. Affinity chromatography of thiamine pyrophosphokinase from rat brain on thiamine monophosphate-agarose. Methods Enzymol 62:105–107, 1979. Y Wakabayashi, A Iwashima, Y Nose. Affinity chromatography of thiamin pyrophosphokinase of rat brain. Biochim Biophys Acta 429:1085–1087, 1976. A AEvarsson, JL Chuang, RM Wynn, S Turley, DT Chuang, WG Hol. Crystal structure of human branched-chain alpha-ketoacid dehydrogenase and the molecular basis of multienzyme complex deficiency in maple syrup urine disease. Structure Fold Des 8:277–291, 2000. F Jordan. Interplay of organic and biological chemistry in understanding coenzyme mechanisms: example of thiamin diphosphate–dependent decarboxylations of 2-oxo acids. FEBS Lett 457:298–301, 1999. C-H Pletcher, E Sax, K Turano, LJ Chang. Effects of structural variations in thiamin its derivatives and analogues. Ann NY Acad Sci 378:454–458, 1982. CJ Oguchi, CJ Sakamoto. Effect of a single, large-dose injection of pyrithiamin on the thiamin deficiency of the mouse. Jpn J Pharmacol 32:775–783, 1982. CJ Machli. Thiamin. In: C.J. Gubler, ed. Handbook of Vitamins. New York: Marcel Dekker, 1991, pp 233–281. G Gubler. Studies on the physiological functions of thiamine I. The effects of thiamine deficiency and thiamine antagonists on the oxidation of alpha-keto acids by rat tissues. J Biol Chem 236:3112–3120, 1961. L Gubler, U Murdock. Effects of treatment with thiamin antagonists, oxythiamin and pyrithiamin and of thiamin excess of the levels and distribution of thiamin in rat tissues. J Nutr Sci Vitaminol (Tokyo) 28:217–224, 1982. S Rindi, LR de Giuseppe, Ventura. Distribution and phosphorylation of oxythiamine in rat tissues. J Nutr 81:47–154, 1963. S Eich, LR Cerecedo. Studies on thiamin analog III. Effects on enzyme systems. J Biol Chem 207:295, 1953.

4 New Perspectives on the Cellular Role of Thiamine Triphosphate and Thiamine Triphosphatase Lucien Bettendorff and Pierre Wins University of Lie`ge, Lie`ge, Belgium

I. INTRODUCTION It is generally assumed that the biologically active form of thiamine (vitamin B1) is thiamine pyrophosphate (or thiamine diphosphate, TDP), a coenzyme that is absolutely required for cellular oxidative metabolism. TDP is generally the predominant form of thiamine (75–90% of total), but free thiamine and two other phosphorylated derivatives—thiamine monophosphate (TMP) and thiamine triphosphate (TTP)-are also present in most tissues. So far, no biological role has been ascribed to free thiamine or TMP. The latter compound might simply be a product of TDP hydrolysis by nonspecific phosphohydrolases. The case of the triphosphorylated derivative, TTP, is more intriguing. In most tissues, it is only a minor compound (0.1–1% of total thiamine). Yet it has been found to be present in all organisms investigated so far, from bacteria to mammals. Thus, although its synthesis presumably costs energy to cells, TTP may be ubiquitous, and we believe that its metabolism and possible biological role deserve further investigation. The history of TTP began 50 years ago, when Rossi-Fanelli et al. (1) reported its presence in rat liver extracts. At about the same time, Kiessling (2) 43

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showed that TTP could be synthesized by bakers’ yeast from thiamine in the presence of glucose. Though it took nearly 30 more years before reliable analytical methods became available for TTP quantification, its presence in many animal tissues, plants, fungi, and bacteria is well established by now (A.F. Makarchikov and L. Bettendorff, manuscript in preparation). From 1969 to 1980, Cooper and associates (see Ref. 3 for review) published data suggesting that TTP played a role in nerve excitability. This idea originated from earlier observations (4–6) that thiamine was released during nerve activity, probably as a result of hydrolysis of thiamine phosphate compounds. However, the evidence implicating TTP remains doubtful, for the methods used to quantify the compound were unreliable until 1980. Thus, the hypothesis of a neuronal-specific role for TTP remains poorly substantiated (7). II. THIAMINE TRIPHOSPHATE, A NEW SUBSTRATE FOR PROTEIN PHOSPHORYLATION? We have shown that TTP activates a high-conductance chloride channel (maxi-Clchannel) in inside-out patches of neuroblastoma cells (8). There was a lag period of several minutes before the channel activated, and the effect of TTP was irreversible, suggesting that a phosphorylation may be involved. Indeed, TTP contains two phosphoanhydride bonds that make it a compound with a high potential for the transfer of phosphoryl groups. To our knowledge, TTP is the only known triphosphorylated compound besides nucleoside 5V-triphosphates. It is thus very appealing to suggest that TTP might be involved in the regulation of some cell activities through protein phosphorylation. Indeed, it was recently shown that TTP phosphorylates a protein, identified as rapsyn, in the electric organ of T. marmorata (9). Rapsyn is essential for the clustering of acetylcholine receptors at the neuromuscular junction, and rapsyn deficiency results in a lethal phenotype in the mouse (10). Protein phosphorylation by TTP was also observed in membranes prepared from rodent brain, but in this case the proteins involved were not identified. In all cases, no protein kinase was added to the preparation, suggesting that either we are dealing with an autophosphorylation or there is an endogenous membrane-associated TTP-dependent protein kinase. The most intriguing finding was that phosphorylation of rapsyn occurred on histidyl residues instead of the usual Ser, Thr, or Tyr residues. Histidine phosphorylation is the predominant signaling system in prokaryotes (11), while no histidine kinases have yet been described in eukaryotes. Though we must be careful before extrapolating from rapsyn to other proteins phosphorylated by TTP, it is possible that the TTP phosphorylation pathway in eukaryotes is a very old

Cellular Role of TTP and TTPase

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mechanism, maybe derived from ancient prokaryotic His-signaling pathways. This view would be compatible with the fact that TTP itself is found in all main taxonomic divisions, including prokaryotes, fungi, and plants (A.F. Makarchikov and L. Bettendorff, manuscript in preparation). An important corollary of this hypothesis would be that, in contrast to what was initially thought, a potential role of TTP would not be limited to neuronal or excitable tissues but would be far more general. III. THIAMINE TRIPHOSPHATE HAS A RELATIVELY HIGH TURNOVER TTP is a minor thiamine compound, accounting for only about 0.1–1% of total thiamine (12), with some exceptions that will be discussed later. However, when cultured neuroblastoma cells were incubated with radioactive thiamine, radioactivity was rapidly incorporated into TTP (Fig. 1). TTP rapidly reached a much higher specific radioactivity than TDP, suggesting that only part of TDP is the precursor for TTP. This observation suggested the existence of two different TDP pools (Fig. 2): one small cytosolic pool of free TDP, which is the precursor of TTP, and one larger cofactor pool bound to apoenzymes (13,14). It can be seen from Figure 1 that the radioactivity is

Figure 1 Specific radioactivities of thiamine derivatives after incubation with 20 AM [14C] thiamine as a function of time. Each point represents the mean F SD for three experiments. The double line represents the maximum specific radioactivities of [14C] thiamine (24 mCi/mmol). (o, thiamine; ., TMP; 5, TDP; n, TTP.) (From Ref. 13.)

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Figure 2 Model for thiamine metabolism in neuroblastoma cells: 1, thiamine pyrophosphokinase; 2, ATP-TDP phosphotransferase (TDP kinase); 3, thiamine triphosphatase; 4, thiamine diphosphatase; 5, thiamine monophosphatase. (From Ref. 13.)

also rapidly incorporated into TMP, though, in animals, TMP cannot be formed from thiamine but is the product of TDP hydrolysis by thiamine diphosphatase (TDPase). These results suggest that, though the main pool of TDP has a low turnover, probably dependent on the turnover of mitochondrial TDP-dependent enzymes, there is a small cytosolic TDP pool that can be rapidly either phosphorylated to TTP or hydrolyzed to TMP. TDPases have been described in the literature, but their specificity has not been proven and no sequence data are available (for review see Ref. 12). IV. THE MECHANISM OF THIAMINE TRIPHOSPHATE SYNTHESIS REMAINS POORLY CHARACTERIZED As originally proposed by Eckert and Mo¨bus in 1964 (15), the synthesis of TTP would be catalyzed by a TDP kinase (TDP:ATP phosphoryltransferase; EC 2.7.4.15) according to the reaction TDP + ATP Z TTP + ADP. Until now, this enzyme has remained poorly characterized. A purification procedure from brewer’s yeast has been described (16), but the specific activity of the purified enzyme was very low. A similar TDP kinase was also partially purified from rat liver (17). On the other hand, Nishino et al. (18) reported the purification of a TDP kinase from beef brain, but in this case the substrate was protein-bound TDP rather than free TDP. Although TTP generally accounts for less than 1% of total thiamine, a few tissues, such as pig skeletal muscle (19), electric organ of Electrophorus electricus (20), and chicken white muscle (21), contain high concentrations of TTP, even more than the cofactor TDP. Kawasaki and coworkers have


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suggested that this high TTP content was correlated with the high amount of adenylate kinase 1 (AK1; EC 2.7.4.3) found in skeletal muscle and electric organ. Indeed, these authors (22) have shown that in vitro AK1 may synthesize TTP according to the reaction TDP + ADP Z TTP + AMP. However, we have recently demonstrated that AK1 knockout mice have normal TTP levels (23), suggesting that in vivo AK1 is not responsible for TTP synthesis even in skeletal muscle. Furthermore, we were able to show that pig and chicken skeletal muscle have similar AK activity as mouse skeletal muscle, but the former have a very low or even absent TTPase activity, suggesting that this enzyme is, at least in part, responsible for the regulation of intracellular TTP concentration.

V. MOLECULAR CHARACTERIZATION OF A SPECIFIC THIAMINE TRIPHOSPHATASE FROM MAMMALIAN BRAIN An important point is that TTP concentrations seem to be highly regulated in brain cells (24,25); i.e., any excess of TTP is rapidly hydrolyzed. Data from Table 1 suggest that this is the case in other tissues as well. We purified a Table 1 TTP Content and TTPase and Adenylate Kinase Activities in Various Tissues

Mouse skeletal muscle wild-type AK1/ Mouse brain wild-type AK1/ Pig skeletal muscle Chicken white muscle E. electricus electric organ a

TTP content (nmol/g of wet weight)

TTPase activity (nmol-g1 of tissue-min1)

Adenylate kinase activity (Amol-mg1-min1)

0.026 F 0.006 0.026 F 0.008

302 F 54 284 F 70

1.3 F 0.2 0.014 F 0.008

F F F F F F F

283 F 57 345 F 62 12.5 F 0.5

0.123 F 0.034 0.011 F 0.007 1.03 F 0.15

NDd

1.6 F 0.1

NDd

—

0.009 0.014 20 18.24 3.2 1.2 3.9

0.003 0.010 2 5.83a 0.4 0.15b 0.5c

According to Ref. 19. According to Ref. 21, assuming that 1 g of muscle contains 171 mg of protein. c From Ref. 20. d ND, not detectable. b

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soluble ThTPase from bovine brain, as previously described (26), and obtained 107 Ag of a homogeneous enzyme prepration from 5 kg of bovine brain (27). The enzyme had to be purified about 45,000-fold before a homogeneous preparation was obtained, suggesting that it is a relatively rare protein in the bovine brain. Mass spectrometry gave a molecular mass of 23,892 Da, while chromatography on Sephadex G-75 yielded a value of 25 kDa, suggesting that the native protein is a monomer, in agreement with previous results (26). The specific activity (Vmax at 37jC) was 9 F 2 Amol-s1-mg1 and Km = 32 F 6 lM. We can thus calculate that the catalytic constant (kcat) and the catalytic efficiency (kcat/Km) are, respectively, 240 s1 and 6 106 s1-M1. In addition to its high catalytic efficiency, an interesting feature of this enzyme is its specificity for TTP; indeed, the purified bovine TTPase did not hydrolyze nucleoside 5V-triphosphates, TDP, TMP or even p-nitrophenylphosphate to any significant extent. The sequences of several internal peptides were obtained by tandem mass spectrometry. Comparison with the known sequences of the GenBank database gave a nearly perfect match with two newly described hypothetical proteins, one in humans called MGC2652 (NM_024328) and one in Macaca fascicularis (AB055296). The human and bovine cDNAs were amplified by RT-PCR using, respectively, human and bovine brain poly(A)+ RNA. They were then cloned in E. coli and sequenced. At the amino acid level, the bovine TTPase has 80% and 79% identity with the human and the macaque enzyme, respectively (Fig. 3A). Analysis of the sequences using the PROSITE motif search revealed the presence of several potential phosphorylation sites present in the three sequences and among them two consensus sites (at positions 34 and 123) for protein kinase C and three consensus sites (at positions 34, 38, and 60) for casein kinase 2. The hydrophobicity plot of the human enzyme is typical of a soluble protein (Fig. 3B), with several highly polar or charged regions. In order to check that the cloned human cDNA indeed encodes a functional TTPase, it was overproduced in E. coli as a GST fusion protein in the presence of IPTG (Fig. 4). Escherichia coli transfected with GST have a relatively low intrinsic TTPase activity (120 F 34 pmol-min1-mg of protein1, n = 7). Actually, bacteria do not appear to contain a specific TTPase but contain nonspecific phosphatases able to hydrolyze ThTP to some extent (Ref. 28; A.F. Makarchikov, unpublished results). The activity was increased over 1000-fold in noninduced GST-TTPase recombinant bacteria, reaching 0.17 Amol-min1-mg1. After induction by IPTG, this activity still increased over 10-fold, reaching 2.1 Amol-min1-mg1 after 4 hours (Fig. 5C). No increase in ThTPase activity was observed after induction in bacteria transfected with GST alone (Fig. 5B). TTP hydrolysis by recombinant GST-


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Figure 3 (A) Alignment by Clustal W of TTPase amino acid sequences deduced from bovine (AF432863), human (AF432862), and macaque (AB055296) cDNA and hydropathy plot of the human enzyme. The sequences corresponding to the three bovine peptides obtained by tandem mass spectrometry are underlined and in bold characters. The cysteyl residues are indicated by an asterisk. (B) Hydrophathy plot of human TTPase by the method of Kyte and Doolittle (Ref. 29). Two particularly charged sequences are indicated. (From Ref. 27.)

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Figure 4 SDS polyacrylamide gel (12%) electrophoresis of E. coli extracts transfected with pGEX containing the sequence of either GST-TTPase or GST alone. The bacteria were grown in 2XYT/ampicillin medium, and overexpression was induced by addition of IPTG (+) 1.5 mg/mL. The arrows indicate the GST protein (25 kDa) or the GST-TTPase fusion protein (50 kDa). (From Ref. 27.)

TTPase resulted in the formation of TDP only. No further hydrolysis of TDP to TMP was observed, as would be the case with unspecific phosphohydrolases such as alkaline or acid phosphatase. Furthermore, when ATP (100 lM) replaced TTP in the incubation medium under the same conditions (Fig. 5D E F), no hydrolysis of ATP was observed. Furthermore, no significant hydrolysis of nucleoside 5’-triphosphates, p-nitrophenyl phosphate, TDP, or TMP by GST-TTPase was observed, suggesting that the recombinant enzyme, like the native TTPase, is highly specific for TTP.

Figure 5 Chromatograms showing specific TTPase activity in E. coli expressing GST and GST-TTPase after a 4-hour incubation with IPTG. The bacteria were lysed in 10% Triton X-100 (30 min in ice), diluted 1000 times in Tris-Cl buffer (20 mM, pH 7.5). The enzyme activity was measured under identical conditions (incubation for 10 min, 37jC), either with 10 lM ThTP (A, B, C) or with 100 lM ATP (D, E, F) as substrate. (A) and (D), control (no enzyme); (B) and (E), extract from E. coli expressing GST induced by IPTG; (C) and (F), extract from E. coli expressing GSTTTPase and induced by IPTG. ATP and ADP were determined by HPLC (30). (From Ref. 27.)


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VI. THE 24-KDA THIAMINE TRIPHOSPHATASE IS HIGHLY EXPRESSED IN MAMMALIAN TISSUES BUT SEEMS TO BE ABSENT IN OTHER ORGANISMS TTPase expression was profiled by dot blot hybridization on an mRNA multiple-tissue-expression array, using the entire cDNA as probe (Table 2). The main conclusion to be drawn from this experiment is that TTPase mRNA is very widely expressed in human tissues but at a low level, in agreement with the high purification factor needed to obtain a homogeneous enzyme preparation from bovine brain. The highest hybridization signal was observed in uterus, testis, and prostate, followed by bladder, kidney, lung, and thyroid gland. Surprisingly, only small signals were found in different brain regions, with no detectable signal in the cerebellum. TTPase mRNA was also poorly expressed in the digestive system, fetal tissues, and transformed human cell lines. No signal was observed in yeast or E. coli. Furthermore, we did not find any significant homology between TTPase and any other known protein. Because the complete genomes of E. coli, S. cerivisiae, C. elegans, D. melanogaster, and A. thaliana are known, this suggests that no homologous enzyme exists in these organisms. This is in agreement with the absence of TTPase activity in the 20- to 40-kDa regions of these organisms as well as in fish and birds (A.F. Makarchikov and L. Bettendorff, unpublished results). Especially in birds and fish, TTPase activity was coeluted with ATPase and p-nitrophenyl phosphatase activity at a higher molecular mass (>80.000 Da), and it cannot be concluded that we are dealing with a specific TTPase in these cases. It must be concluded that, so far, a specific TTPase could be characterized only in mammals.

VII. CONCLUSIONS TTP exists in all organism studied so far: animals, plants, fungi, and prokaroytes. It may phosphorylate rapsyn in T. marmorata electric organ as well as unidentified proteins in rodent brain (9). This phosphorylation mechanism may be part of a new signaling cascade, the significance of which has still to be established. The only physiological effect of TTP demonstrated up to now is the activation of large unitary conductance Clchannels in the plasma membrane of neuroblastoma cells (8). In this case a phosphorylation of the channel was suggested. TTP probably appeared early during evolution, as documented by its existence in bacteria. TTP phosphorylates a His residue in rapsyn, but we do not know yet if this can be related to ancient His phosphorylation pathways in prokaroytes.


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Table 2 Dot Blot Analysis of the mRNA Distribution of TTPase in Human Tissues Using a Human Multiple-Tissue-Expression Array (Clontech) Tissue Nervous system Whole brain Cerebral cortex Frontal lobe Parietal lobe Occipital lobe Temporal lobe P.G.b of cerebral cortex Pons Cerebellum left Cerebellum right Corpus callosum Amygdala Caudate nucleus Hippocampus Medulla oblongata Putamen Nucleus accumbens Thalamus Tranformed cell lines Leukemia, HL-60 HeLa Leukemia, K562 Leukemia, MOLT-4 Burkitt’s lymphoma, Raji Burkitt’s lymphoma, Daudi Colorectal adeno carcinoma Lung carcinoma, A549

Score

Tissue

Score

Cardiovascular tissue NDa + + ++ ++ + + + ND ND + + + + + + + +

Score

Reproductive and urinary tracts

Heart Aorta Atrium, left Atrium, right Ventricle, left Ventricle, right

+ + + + + ++

Placenta Uterus Ovary Prostate Testis Bladder

+ ++++ ND ++++ ++++ +++

Interventricular septum Apex of the heart Lung Trachea

++

Kidney

+++

FETAL TISSUE Brain Heart Kidney Liver Spleen Thymus Lung

+ ND + + + ++ +

OTHER Thyroid gland Adrenal gland Skeletal muscle Liver

+++ ++ ++ ++

+ +++ +

ND + ND + +

DIGESTIVE SYSTEM Esophagus + Stomach + Duodenum + Jejunum ++ Ileum ND Ilocecum ND Appendix ND Colon, ascending ND Colon, transverse + Colon descending + Rectum + Pancreas +

+

Salivary gland

++

+ +

Tissue

IMMUNE SYSTEM Spleen

+

Thymus

++

Yeast

ND

Leucocyte, peripheral

+

E. coli

ND

Bone marrow

++

A single + indicates that the signal was just detectable with the naked eye. Each additional + corresponds to approximately a doubling of the intensity of the signal. a ND, not detectable. b Paracentral gyrus. Source: Ref. 27.

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Figure 6 Hypothesis illustrating the role of TTP in mammalian cells. Upon a signal there is either down-regulation of TTPase or an up-regulation of TDP kinase, leading to increased steady-state concentrations of TTP in the cells. TTP can then phosphorylate specific protein targets and induce a response. An example of such a mechanism might be the phosphorylation of rapsyn, leading to the clustering of acetylcholine receptors at the neuromuscular junction under the influence of agrin.

In most mammalian tissues, and in brain in particular, the intracellular TTP concentration appears to be tightly regulated. The virtually absolute specificity of the 25-kDa TTPase as well as its high catalytic efficiency suggest that this enzyme is responsible for the regulation of intracellular TTP concentrations at a very low level (often 600

2650 F 210 685 F 70 0.60 F 0.08 >2.5 >5 0.07 F 0.01 294 F 20

397 F 20 530 F 45 0.13 F 0.01 > >5 > >25 > >7 105 F 6

265 150 1.2 0.051 0.25 0.09 105

150 100 0.10 0.05 0.24 0.04 45

5 4 0.004 0.002 0.03 0.003 2

F F F F F F F

13 14 0.2 0.002 0.03 0.01 6

Rate-limiting steps are indicated in boldface type. H/D exchange rate constants (kobs) were determined according to Ref. 3 at 5jC. b Rate constants of covalent pyruvate binding to TDP (k2), decarboxylation of LTDP (k3), and acetaldehyde release from HETDP (k4) at 30jC in 50 mM sodium phosphate, pH 6.0. a

Figure 2 Active site of ZmPDC. The crystal structure of active-site residues of ZmPDC with the enzyme-bound cofactor TDP in its typical V-conformation and a water molecule. (From Ref. 10.)

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volved in decarboxylation as well as in substrate binding, we assume this residue to be uncharged, because otherwise charge repulsion with the negatively charged carboxylate of pyruvate/LTDP would complicate a proper binding of pyruvate and the stereochemical control of the decarboxylation of LTDP. In accordance with theoretical studies on LTDP decarboxylation (17), our experimental data demonstrate that a perpendicular conformation of the carboxylate of LTDP is mandatory for a fast decarboxylation. Once the crucial interaction between LTDP and the protein is removed by methylene-shortening of the interacting sidechain (Glu!Asp), the rate of decarboxylation of LTDP is diminished. The methylene approach was again successfully applied by engineering an Asp27Glu variant. The sole occurrence of HETDP in steady state points to a rate-limiting product elimination from HETDP (Fig. 1B). At first sight, either impaired protonation of the a-carbanion/enamine form of HETDP or deprotonation of the Ca-hydroxyl function of HETDP might be affected. But intriguingly, the evidenced ability of Asp27Glu and other Asp27 variants (18) to catalyze acetoin formation clearly demonstrates the presence of the acarbanion/enamine, which is, from a chemical point of view, expected to be far more reactive than its protonated counterpart. A similiar deficiency in catalyzing acetaldehyde elimination from HETDP is observable in the His113Lys variant (Fig. 1B). Since His113 is in hydrogen-bonding distance to Asp27, both residues are apparently forming a functional dyad that is responsible for the protonation of the a-carbanion/ enamine of HETDP. Perturbation of this PDC-specific proton relay system in either geometrical or thermodynamical terms deminishes the catalytic ability to protonate HETDP by several orders of magnitude (Table 1). It is still an open question whether the solvent molecule found in striking distance to Asp27 in the X-ray structure or another one serves as the orginal proton donor for charging this relay (Fig. 2). When catalysis is allowed to proceed in D2O, both pyruvate binding to TDP as well as the product release show primary isotope effects, whereas decarboxylation of LTDP is nearly unaffected. Aside from the Glu473, Asp27, and His113 variants, which are in close proximity to the thiazolium moiety of TDP, we investigated the intermediate distribution of the Glu50Gln variant. This glutamate side chain is in hydrogen-bonding distance to N1V of the aminopyrimidinium ring of TDP, and this short distance renders Glu50 a chemical trigger for the reactivity of the 4Vamino group of TDP. The C2-H ionization rate in Glu50Gln is decreased in comparison to the wild-type enzyme, albeit to a lesser extent than found in homologous variants of other TDP-dependent enzymes, and is still far from being rate limiting (Table 1). The steady-state distribution of TDP-derived intermediates in this down-tuned ‘‘4V-amino variant’’ apparently consists of TDP and HETDP (Fig. 1B), demonstrating the involvement of the 4V-amino

How Thiamine Works in Enzymes

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group in substrate binding as well as in the cleavage of HETDP to TDP and acetaldehyde. Given the essential interaction between Glu473 and the carboxylate group of LTDP and the resultant conformation of this tetrahedral intermediate, the 4V-amino group is in a favorable positon to catalyze protonation of the Ca-alkoxidic form of LTDP. The invisibility of LTDP in steady state in Glu50Gln points to a concerted action of nucleophilic attack of the C2 ylide of TDP on the keto group of pyruvate and protonation of the so-formed intermediate, rather than a stepwise catalysis. The conserved spatial proximity of the 4V-amino group to the a-hydroxyl group of HETDP after decarboxylation of LTDP very likely promotes deprotonation of the ahydroxyl group by the 4V-amino (imino) group in the course of product elimination. Based on the steady-state distribution of the wild-type enzyme and the engineered active-site variants with defined deficiencies (Table 1) in C2-H ionization (Glu50Gln), covalent binding of pyruvate (Glu50Gln, Glu473Asp), decarboxylation of LTDP (Glu473Asp), and acetaldehyde elimination from HETDP (Glu50Gln, Asp27Glu, His113Lys), a mechanistic model of the catalytic cycle can be suggested (Fig. 3). 1. C2-H Ionization of Enzyme-Bound TDP In accordance with our earlier results (3), the C2-H deprotonation is catalyzed by the 4V-imino group of TDP since a perturbation of the proton relay involving the conserved glutamate and N1V of ThDP as well as the 4V-amino group of the cofactor results in decreased deprotonation rates in PDC, whereas the exchange of all other potential acid/base catalysts in the active site does not impair this initial catalytic step (Table 1). 2. Covalent Substrate Binding The C2 ylide nucleophilically attacks the Ca of the substrate, which is orientated by the interaction of Glu473 with its carboxylate group. This interaction can provide not only for steric orientation but also for increased electrophilicity of the substrate. The protonation of the Ca-oxygen of the substrate, catalyzed by the 4V-immonium group of ThDP, appears to proceed in a concerted manner with covalent bond formation, rather than in a stepwise manner. 3. Decarboxylation of LTDP The stereochemical control of the decarboxylation of LTDP by the enzyme component is one of the most exciting findings in these studies. Apparently, the interaction between Glu473 and the carboxylate of LTDP leads to a perpendicular orientation of the LTDP-carboxylate, making carbon dioxide release easier. Engineering a ‘‘missing interaction’’ Glu473Asp variant results

Figure 3 Suggested model of catalysis. Proposed catalytic mechanism of PDC. The suggested roles of active-site residues for different steps of catalysis are consistent with the intermediate distribution during enzymic decarboxylation of pyruvate by PDC and its variants. This mechanism includes the action of two independent proton relay systems catalyzing the activation of TDP (Glu50N1V-4V-NH2, step 1), substrate binding (Glu50-N1V-4V-NH2, step 2), and acetaldehyde release (Glu50-N1V-4V-NH2 and His113-Asp27, step 4) as well as the stereochemical control of decarboxylation by glutamate 473, inducing a perpendicular orientation of the substrate carboxylate to the thiazolium ring of the enzyme-bound TDP (Glu473, steps 2 and 3), shown in the Newman projection.

66 Tittmann et al.


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in a 3000-fold slower decarboxylation of LTDP in this variant (k = 0.13 s1). Still, the rate of decarboxylation in Glu473Asp is far above those found for the nonenzymic system under comparable conditons (k = 5 105 s1; Ref. 19). This might be due to a less polar environment in the active site, as already shown to promote a large acceleration of the decarboxylation in model systems (20). 4. Acetaldehyde Release from HETDP The elimination of acetaldehyde from the a-carbanion/enamine of HETDP is catalyzed in a concerted action of a second proton relay spanning His113, Asp27, and an active-site water and the 4V-iminogroup of the cofactor involving protonation of the Ca of HETDP as well as deprotonation of the a-hydroxyl function of this intermediate. As already outlined, the conformational orientations of the covalent intermediates as well as the reported ability of Asp27 variants to catalyze acetoin/acetolactate formation clearly favor our proposed mechanism comprising carbanion protonation by the (His113/ Asp27/H2O)-relay and deprotonation of the hydroxyl function by the 4Vimino group of the enzyme-bound cofactor. B. Acetohydroxyacid Synthases The class of the TDP-dependent acetohydroxyacid synthases (AHAS, EC 4.1.3.18) occurs in plants and bacteria and catalyzes the condensation of onemolecule pyruvate with either another pyruvate or a-ketobutyrate to yield the amino acid precursors acetolactate and acetohydroxybutyrate, respectively. Essentially, the chemical path of the first half reaction of catalysis culminating in the buildup of the a-carbanionic form of HETDP is identical to PDC (compare Fig. 3) and all other pyruvate converting TDP-dependent enzymes. The chemical fate of the latter intermediate in AHAS, however, differs from all other enzymes, being subject to covalent addition of a second substrate molecule followed by the liberation of the respective acetohydroxyacid. Apparently, the catalytic cycle of AHAS includes a covalent TDPadduct, in addition to LTDP, HETDP, and TDP. Since all chemical attempts to synthesize the covalent acetolactyl-TDP (ALTDP) intermediate de novo failed to date, we investigated the intermediate distribution of the Asp28Ala ScPDC variant, which is the only non-AHAS TDP-dependent enzyme known so far that preferentially forms acetolactate instead of any other product, even though the overall catalytic constant is very small (21). NMR-based investigation of the intermediate distribution of this variant at steady state indeed revealed the predominant presence of an acid-stable covalent TDP adduct, corresponding to neither LTDP, HETDP, nor TDP. The 1H NMR spectrum of this intermediate is composed of all resonances typical of a covalent TDP

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adduct, with slightly altered chemical shifts and two additional methyl singlets in comparison to TDP, thus giving rise to the reasonable conclusion that this intermediate is the covalent ALTDP adduct formed on addition of pyruvate to HETDP (Fig. 4A). Using this enzymatically generated TDP adduct and the chemically synthesized LTDP and HETDP as standards, we investigated the intermediate distribution of AHAS II from E. coli in steady state (Fig. 4A). As depicted in Figure 4, the most populated intermediate in EcAHAS II is TDP, whereas only small amounts of LTDP, HETDP, and ALTDP are detectable and even those only at very high concentrations of AHAS (c = 30 mg/mL). As already outlined for the intermediate analysis in PDC, a set of experiments was conducted varying the reaction times of AHAS with the substrate and with different pyruvate concentrations. The intermediate distributions are the same for pyruvate concentrations between 25 and 100 mM and varied mixing times (0.1–1 s), which clearly point to intra(uni-)molecular covalent addition of the first substrate to TDP and the second one to HETDP. In a manner analogous to the transformation of the quantitative distribution of the intermediates in PDC into microscopic rate constants, we determined those for AHAS, assuming the catalytic cycle to comprise (1) intramolecular addition of the first pyruvate, (2) decarboxylation of LTDP, (3) intramolecular addition of the second pyruvate to HETDP and, finally, (4) liberation of acetolactate (Table 2). Evidently, the addition of pyruvate to the C2 of TDP is nearly totally rate limiting, whereas all other catalytic steps are comparatively fast. The intermediate analysis of AHAS in steady state in the presence of pyruvate and a-ketobutyrate (data not shown) could only reveal detectable populations of TDP and LTDP. This solves the puzzle as to why AHAS II prefers a-ketobutyrate 60-fold over pyruvate as the second substrate, though kcat is nearly identical for acetolactate and acetohydroxybutyrate formation (22). Since the kinetic significance of the covalent additon of the second substrate regarding kcat is very low, the intermediate partitioning of HETDP is reflected only in the ratio of the different products but not in kcat itself. Consequently, a-ketobutyrate competes favorably with pyruvate as the electrophile toward the a-carbanion of HEThDP. The recently solved X-ray crystallographic structure of AHAS (see Chapter 15) will enable the rational design of active-site variants, probably showing significant perturbations and deficiencies in single steps of catalysis. Determination of the intermediate distributions in these perturbation variants will reveal mechanistic principles of catalyis at a molecular level. C. Pyruvate Oxidase Pyruvate oxidase (POX, EC 1.2.3.3) is an FAD and TDP-dependent enzyme catalyzing the oxidative decarboxylation of pyruvate, yielding acetylphos-

Figure 4 Detection of covalent intermediates in the superfamily of TDP-dependent enzymes, including acetohydroxyacid synthase (AHAS), pyruvate oxidase (POX), and transketolase (TK). (A) Covalent intermediates at steady state of EcAHAS II and of Asp28Ala PDC from Saccharomyces cerevisiae, which produces acetolactate exclusively. (B) Covalent intermediates at steady state of LpPOX in the presence and absence of phosphate and after complete reduction of the enzyme-bound FAD. (C) Covalent intermediates during conversion of different donor substrates (h-hydroxy-pyruvate, xylulose 5-phosphate, and fructose 6phosphate) by TK. The intermediates I1 and I2 can be attributed to the 2-[2-(1,2,3,4,5-pentahydroxy)-pentyl]-TDP and 2-[2(1,2,3,4,5,6-hexahydroxy)-hexyl]-TDP derivatives based on the chemical pathway. (From Ref. 21.)

How Thiamine Works in Enzymes 69

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Table 2 Microscopic Rate Constants of Elementary Catalytic Steps of EcAHAS II Wild Type in the Presence of Pyruvate (P + P, Product Acetolactate) or Pyruvate and aKetobutyrate (P + KB, Product Acetohydroxybutyrate)

EcAHAS II (P + P) EcAHAS II (P + KB) ScPDC Asp28Ala (P + P)

kcat (s1)

k2 (s1) C–C bonding TDP + pyruvate

k3 (s1) CO2 release

k4 (s1) C–C bonding HETDP + 2nd substrate

k5 (s1) product release

20 F 0.2 20 F 0.2 0.052 F 0.001

24 F 4 21 F 4 >0.4

530 F 70 399 F 54 > >1

1060 F 175 >2000 0.36 F 0.05

176 F 36 >2000 0.073 F 0.007

Rate constants of covalent pyruvate binding to TDP (k2), decarboxylation of LTDP (k3), covalent binding of the second substrate molecule (k4), and product release (k5) were determined at 37jC in 50 mM potassium phosphate, pH 7.6 (EcAHAS) and in 50 mM potassium phosphate, pH 6.0 at 30jC (ScPDC), respectively. Rate-limiting steps are indicated in boldface type.

phate or acetate and carbon dioxide. As in PDC and AHAS, the first halfreaction at TDP leads to the formation of the a-carbanion/enamine of HETDP. Again, this key intermediate is the branching point in catalysis. Rather than releasing acetaldehyde (PDC) or condensing with another substrate molecule (AHAS), HETDP in POX transfers two electrons to the nearby FAD and is oxidized to AcTDP. Once the electron transfer between HETDP and FAD is blocked either by completely reducing the enzyme-bound FAD prior to or in the course of catalysis or by replacing FAD with 5-deaza-FAD,* all active sites in POX are indeed quantitatively occupied by HETDP, as evidenced by 1H NMR analysis of the isolated covalent TDP intermediates (Fig. 4B). These observations hold true for the POX from both Lactobacillus plantarum (LpPOX) and Escherichia coli (EcPOX). Furthermore, the accummulation of AcTDP in LpPOX in the absence of phosphate, which was indirectly inferred by earlier kinetic studies (24), could now be directly detected. In contrast, LTDP and a minor fraction of HETDP are detectable at steady state in the presence of Pi (Fig. 4B). Surprisingly, no AcTDP is detectable under those conditions, even though the phosphorolytic decomposition of AcThDP was presumed to be partially rate limiting (24). It must be noted, however, that the 1H NMR chemical shifts and the stability of the tetrahedral adduct between AcThDP and Pi are still unknown. Consistent with this, a concerted action of an intramolecular redox reaction and a nucleophilic attack of Pi on AcTDP could explain the absence of AcTDP. * 5-Deaza-FAD in POX was already shown not to accept electrons in the course of catalysis (23).


71

In conclusion, it is now possible to observe, directly and independently, intermediates in POX at both TDP and FAD using 1H NMR and FAD absorbance. A combination of these approaches to studying rationally designed perturbation variants (active-site side chains, cofactor analogs) will provide important insights into the principles of biological electron transfer between the two cofactors as well as catalytic events at TDP. D. Pyruvate Dehydrogenase Pyruvate dehydrogenase (PDH, EC 1.2.4.1) is a key metabolic enzyme in many species linking glycolysis and citrate cycle. This multienzyme complex consists of three components (E1, E2, E3) and catalyzes the oxidative decarboxylation of pyruvate, yielding the essential metabolite acetyl-CoA. The ThDP-dependent E1 component (decarboxylase component) binds and decarboxylates pyruvate, resulting in the formation of the a-carbanion/enamine of HETDP. Within the framework of our 1H NMR studies we investigated kinetically the formation of LTDP and HETDP at E1 under single-turnover conditions for mammalian E1 only, as well as for the entire PDH complex. As expected, all active sites in both E1 only and the complex are occupied by HETDP after completion of the single-turnover reaction (data not shown). The microscopic rate constants of substrate binding (kC–C) and decarboxylation (kdec), however, are strikingly different for the isolated E1 component and the complex. Most remarkably, covalent pyruvate binding at C2 of TDP is composed of two phases and is slower in E1 only (kC–C1 = 2 s1, kC–C2 = 0.01 s1) than in the PDH complex (kC–C = 102 s1), whereas decarboxylation of LTDP proceeds with comparable rates of kdec (E1) = 2 s1 and kdec (complex) = 6 s1. As already demonstrated for the other TDP-dependent enzymes, it is now also possible to directly observe covalent intermediates and to kinetically resolve their interconversions at E1 in the complex or alone. This feasibility is the prerequisite to investigate at a molecular level the mechanism of catalytic steps and principles of the regulatory control of this important enzyme. E. Transketolase Transketolase (TK, EC 2.2.1.1) plays a vital role in the pentose phosphate shunt of the glycolytic pathway in all organisms. This enzyme catalyzes the ketol transfer of 2-carbon fragments between ketose phosphates (donor substrate) and aldose phosphates (acceptor substrate). Donor substrate conversion in TK can be followed directly by 1H NMR analysis of the isolated covalent TDP intermediates. Addition of the native

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donor substrates D-xylulose 5-P and D-fructose 6-P to TK leads to the transient formation of acid-stable, covalent TDP intermediates (I1 and I2, respectively), both decomposing to DHETDP (Fig. 4C). In the further progress of the reaction, DHETDP is converted to TDP (data not shown). Even though the 1H NMR signals of the sugar moieties of I1 and I2 interfere with the resonances of the donor sugar substrates and therefore could not be assigned unequivocally, the chemical pathway suggests I1 and I2 are nothing else but the 2-[2-(1,2,3,4,5-pentahydroxy)-pentyl]-TDP and 2-[2-(1,2,3,4,5,6hexahydroxy)-hexyl]-TDP derivatives, respectively. In accordance with very recent X-ray crystallographic data (25), conversion of the artificial donor substrate h-hydroxypyruvate leads to the accumulation of DHETDP in the course of catalysis in TK (Fig. 4C). Contrary to this artificial ketose surrogate, the key metabolite pyruvate is not converted by TK. NMR analysis of the isolated intermediates derived on addition of pyruvate to TK revealed the exclusive presence of TDP and the absence of LTDP and HETDP (data not shown). Conclusively, carbon– carbon bonding between pyruvate and TDP is blocked in TK. Hence, the cytosolic sugar-converting TK is not inhibited by pyruvate-derived, slow interconverting or stable covalent intermediates, thus excluding a fatal blockage of the sugar metabolism. These data impressively demonstrate that the interaction of just one hydroxyl group of the substrate with the protein may account for absolute substrate-type specificity, thereby avoiding interference with other metabolic pathways. As outlined earlier, the newly developed ability to directly observe catalysis in TDP-dependent enzymes using 1H NMR can be successfully applied not only to enzymes that act on pyruvate but also to a prominent enzyme in sugar metabolism. The forthcoming correlation of X-ray crystallography (see Chapter 11), NMR spectroscopy, and protein engineering will combine structural and functional aspects of catalysis in TK. IV. SUMMARY AND CONCLUSIONS Thiamine catalysis in enzymes proceeds via covalent intermediates at C2 of the enzyme-bound TDP. The occurrence of the key intermediates LTDP, HETDP, AcTDP, and DHETDP in enzymic catalysis was directly demonstrated by 1H NMR for several TDP-dependent enzymes, including pyruvate decarboxylase, acetohydroxyacid synthase, pyruvate oxidase, pyruvate dehydrogenase, and transketolase, some of those intermediates (such as LTDP) for the very first time. In addition to the validation of the different chemical pathways of the investigated enzymes, the quantitative distribution of these covalent intermediates under steady-state or single-turnover conditions provides access to the microscopic rate constants of single steps in catalysis, invoking covalent bond formation or cleavage. The possibility of chemically


73

freezing an enzyme in action and directly analyzing the distribution of all covalent intermediates is rare in enzymology. This ‘‘one snapshot—all covalent intermediates’’ approach features enormous advantages in observing and understanding the mechanism of a protein. Whereas X-ray crystallographic snapshots account for essential structural information of intermediates, those studies are restricted to enzymic systems with one stable intermediate. Now these limitations in terms of a profound kinetic and thermodynamic characterization at an atomic level can be overcome by the combination of rapid quenched flow and subsequent quantitative 1H NMR spectroscopic analysis of the isolated intermediates. Essentially, the intermediate distribution and consequently the respective microscopic rate constants can be determined independent of the magnitude of k cat or instrumental limitations (dead time) in the steady-state approach. Therefore, rate constants are measurable that may range up to thousands per second. Given the fact that thiamine catalysis in most cases involves irreversible steps (decarboxylation of LTDP), relaxation methods are not suitable for observing substrate turnover in these enzymes. In addition to enabling the direct and parallel detection of all key intermediates in enzymic thiamine catalysis and the kinetic analysis of the wild-type enzymes, this method is very valuable for a comprehensive elucidation of mechanistic principles of enzymic thiamine catalysis. As exemplified with pyruvate decarboxylase, we could prove the specific involvement of active-site side chains in distinct catalytic steps by investigating the intermediate distribution of rationally designed perturbation variants. These results suggest a model that comprises the concerted display of two independent proton relays and an acidic residue. The first proton relay spanning the g-carboxylate of a conserved glutamate (Glu50 in ZmPDC), N1V and 4Vamino group of the enzyme-bound cofactor accounts for sufficiently fast deprotonation at C2 of TDP and the protonation of the alkoxidic form of LTDP after covalent binding of pyruvate. The decarboxylation of this first covalent intermediate (LTDP) is stereochemically controlled by the perpendicular orientation of the negatively charged carboxylate of LTDP, very likely due to a hydrogen bond between Glu473 and LTDP and resulting in a maximal resonance stabilization of the resulting a-carbanion by orbital overlap with the thiazolium ring. The release of acetaldehyde from the a-carbanion/ enamine of HETDP requires the concerted action of two converging proton relays. Whereas the catalytic dyad of His113 and Asp27 leads to the activation of an active-site bound water that decomposes to OH and very likely transfers one proton to HETDP, the aminopyrimidinium relay of TDP catalyzes the deprotonation of the a-hydroxyl function of HETDP (see mechanism in Fig. 3). Going beyond the class of pyruvate decarboxylase, it will be exciting and instructive to prove whether some of the mechanistic principles suggested

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for PDC catalysis hold also true for other TDP-dependent enzymes, especially for those catalytic steps leading to the buildup of the a-carbanion/enamine of HETDP as the key intermediate of all pyruvate-converting TDP-dependent enzymes and the central branching point in enzymic thiamine catalysis. It has already been shown consistently for most of these enzymes that the adopted V-conformation of the cofactor in the active site as well as the functional triad of a conserved glutamate, N1V nitrogen and 4V-amino group of the cofactor itself are crucial for its activation with respect to C2-H ionization. But it still remains to be clarified in which way the different enzymic environments are orienting the substrate molecule in the course of the initial covalent bond formation between TDP and pyruvate and which interactions account for this correct orientation. A further major issue is to prove whether the decarboxylation of LTDP is stereochemically controlled in other TDP-dependent enzymes as well, or whether other mechanisms, such as environmental control, account for proficient and fast carbon dioxide release. Although similar principles might govern the first half reaction of pyruvate converting enzymes, culminating in the formation of the a-carbanion/enamine of HETDP, all subsequent steps are essentially divergent for those enzymes. The forthcoming and challenging task will be to dissect the molecular determination of the different chemical fates of this key intermediate in enzymic thiamine catalysis. The combination of X-ray crystallography– based structural biology, the newly established intermediate NMR approach accounting for dynamic and mechanistic aspects of thiamine catalysis, and a rational protein engineering will help to extend and evolve our understanding of why the a-carbanion/enamine of HETDP/DHETDP is subject to either covalent ligation of a second substrate (acetohydroxyacid synthase, transketolase) or intramolecular redox processes and acyl transfer (pyruvate dehydrogenase, pyruvate oxidase). The molecular elucidation of the highly sophisticated catalytic machinery in pyruvate decarboxylase involving a decarboxylase-specific proton relay for the protonation of HETDP (enamine/a-carbanion) might therefore be seen as a starting point for a comprehensive analysis of the divergent and convergent catalytic principles in enzymic thiamine catalysis at an atomic level. ACKNOWLEDGMENTS We gratefully acknowledge helpful discussions with P. Frey, R. L. Schowen, D. Kern, and S. Ghisla and the latter for providing a sample of 5-deazaFAD. We thank J. Brauer, C. Simm, and B. Seliger for technical assistance. We thank M. Vyazmensky and A. Bar-Ilan for preparation of the plasmid used to express EcAHAS II and A. Chang, C.-Y. Huang, and Y.-G. Wu for providing the various ZmPDC mutant plasmids. This work was supported


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by the Deutsche Forschungsgemeinschaft and the Fonds der chemischen Industrie, Australian Research Council grant number A09800834, U.S. grant NIH-GM-50380, the Swedish Science Research Council, and a seed grant from the Research and Development Authority of Ben-Gurion University. REFERENCES 1. 2. 3.

4.

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6 Thiamine-Dependent Enzymes as Catalysts of C–C Bond-Forming Reactions: The Role of ‘‘ ‘‘Orphan’’ ’’ Enzymes Michael Mu¨ller and Georg A. Sprenger* Forschungszentrum Ju¨lich GmbH (Research Centre Ju¨lich), Ju¨lich, Germany

I. INTRODUCTION Thiamine diphosphate (TDP)-dependent enzymes are able to cleave or form carbon–carbon (C–C) bonds. Several TDP enzymes have already been used as catalysts in chemoenzymatic syntheses, making use of their carboligating potential, e.g., phenylacetylcarbinol formation by pyruvate decarboxylase (PDC) or formation of rare sugars by transketolases (TK) from various organisms. For recent reviews, see Refs. 1–6. The purpose of the present chapter is to review the lesser-known TDP enzymes (‘‘orphan enzymes’’) as they have potential to catalyze fascinating C–C bond-forming reactions. Some of the enzymes we are going to discuss have only recently been described. In some cases, only a respective TDP-dependent activity has been detected in crude extracts. Therefore, closer inspection of their substrate ranges and potential side reactions (e.g., acyloin condensations) is warranted.

*Current affiliation: Universita¨t Stuttgart, Stuttgart, Germany.

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II. ACYLOIN CONDENSATIONS Various TDP-dependent a-keto acid decarboxylases have been described as catalyzing C–C bond formation and/or cleavage. For example, pyruvate decarboxylase (PDC) catalyzes as a main reaction the nonoxidative decarboxylation of pyruvate. In a side reaction, an activated acetaldehyde is ligated with benzaldehyde in a benzoin-condensation-like manner (‘‘acyloin condensation’’) to form (R)-phenylacetyl carbinol [(R)-PAC], the precursor of ()-ephedrine (for recent reviews see Refs. 1 and 3–5). The potential of benzoylformate decarboxylase (BFD) to catalyze C–C bond formation was first reported by Wilcocks et al. using crude extracts of Pseudomonas putida (7,8). They observed the formation of (S)-2-hydroxy1-phenyl-propanone [(S)-2-HPP] when benzoylformate was decarboxylated in the presence of acetaldehyde. Advantageously, aldehydes can be used directly without prior decarboxylation of the corresponding and more expensive a-keto acids. And BFD is able to bind a broad range of different aromatic and heteroaromatic aldehydes to TDP prior to ligation to acetaldehyde (9). Acetohydroxyacid synthase (AHAS) has long been known for the synthesis of (S)-acetolactate starting from two molecules of pyruvate (10,11). Most recently, Sergienko and Jordan described the same reaction catalyzed by a variant of PDC (12). Corresponding to this, Chipman and coworkers published the AHAS-catalyzed reaction of pyruvate with benzaldehyde to form (R)-PAC (13). Recently, asymmetric acyloin condensation catalyzed by phenylpyruvate decarboxylase was described by Patel et al. (14,15). Extensive work with regard to asymmetric C–C bond formation has been conducted on transketolase (TK) from different sources and recently reviewed (2,4,6); one of the substrates TK can use is hydroxypyruvate, which thereby undergoes decarboxylation. The transketolase-related 1-deoxy-D-xylulose-5-phosphate synthase (DXS) from Escherichia coli has already been used for the synthesis of deoxysugars in isotope-labeled or unlabeled form using pyruvate as donor (16–19). From this it is obvious that TDP-dependent a-keto acid decarboxylases are able to catalyze a diverse set of asymmetric C–C bond-forming reactions. Here we want to draw attention to some lesser-known TDP-dependent enzymes. For example, TDP-dependent benzaldehyde lyase (BAL) from Pseudomonas fluorescens, which as the wild-type enzyme does not catalyze decarboxylation of a-keto acids, has been identified as a potent catalyst for asymmetric C–C bond formation or kinetic racemic resolution via C–C bond cleavage (20,21). A detailed description on the use of both benzoylformate decarboxylase and benzaldehyde lyase can be found in Chapter 7.

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III. RANGE OF REACTIONS CATALYZED BY ‘‘ORPHAN’’ TDP DECARBOXYLASES The use of C–C bond-forming TDP-dependent decarboxylases PDC from various microorganisms, BFD from Pseudomonas species, and phenylpyruvate decarboxylase has recently been surveyed (3–5) and therefore will not be dealt with here in much detail. Other TDP-dependent decarboxylation reactions are given in Scheme 1 and will now be discussed. Indolepyruvate decarboxylases (IPDC) and variants thereof might be another class of enzymes eligible to enlarge the substrate spectra amenable to asymmetric C–C bond formation [Scheme 1, Eq. (1)]. The structure and function of these IPDC enzymes have been reviewed by Koga (22). Indole-3pyruvic acid is the product of deamination of L-trytophan by a transaminase. The role of IPDC is the decarboxylation to indole-3-acetaldehyde, which is then converted into indole-3-acetic acid in a variety of organisms, e.g., Enterobacter cloacae and Erwinia herbicola (22,23), and also in plants, where it serves as a phytohormone that plays a central role in plant growth and development. The gene for IPDC from E. cloacae was cloned, and its derived protein sequence showed a high degree of similarity with the PDC sequences especially from Saccharomyces cerevisiae and to a lesser degree with the PDC from Zymomonas mobilis (22). So it came as no surprise that IPDC also decarboxylates pyruvic acid (Km of 2.5 mM) besides its cognate substrate, indole-3-pyruvic acid (Km of 15 lM). Active IPDC from E. cloacae forms a homotetramer as PDC enzymes of S. cerevisiae and Z. mobilis. Whether IPDC serves an acyloin condensation function is unknown but deserves further observation because the indole side group opens an interesting product alternative. The lactic acid bacterium Lactococcus lactis IFPL730 produces high amounts of the volatile methional [Scheme 1, Eq. (2)], which is a component of good cheddar cheese flavor and also a potent odorant in Camembert cheese. Methionine is a proteolysis product in milk (mainly from the main milk protein casein) and is converted to the a-ketoacid 4-methylthio-2ketobutanoate (KMBA) by transamination. In cheese KMBA either can be converted to 2-hydroxy-4-methylthiobutyrate or is decarboxylated to methional. Recently, a TDP-dependent activity was described that occurred in crude extracts of L. lactis. This decarboxylation step is catalyzed by the novel 4methylthio 2-ketobutanoate decarboxylase (24). The enzyme activity of this ketoacid decarboxylase was partially purified to allow initial characterization; a major band of 30 kDa was enriched, but further purification seems to be necessary. Interestingly, the highest relative activity was with a-ketoisovalerate [100%; Scheme 1, Eq. (3)], then a-ketoisocaproate (74%), a-ketomethyl-valerate (40%), KMBA (20%), and phenylpyruvate (11%). The

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Scheme 1 Different types of reactions catalyzed by TDP-dependent a-ketoacid decarboxylases.


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specific activity for KMBA was 5 mU/mg of protein, with a Km of about 1 mM. A possible acyloin condensation reaction, however, was not analyzed in this first publication (24). Sulfopyruvate decarboxylase [Scheme 1, Eq. (4)] is an extremely oxygensensitive enzyme from methanobacteria such as Methanococcus jannaschii. It plays a role in the biogenesis of coenzyme M (2-mercaptoethanesulfonic acid), which is one of several coenzymes involved in the formation of methane in the methanobacteria. The genes comD and comE were recently identified (25). The gene products, ComD and ComE, form a heteromeric TDP enzyme, sulfopyruvate decarboxylase, with a likely structure of an a6h6-dodecamer. The ComE subunit contains the TDP-binding motif, whereas ComD displays a similarity to the pyruvate-binding domain of related enzymes like acetolactate synthases, pyruvate oxidases, and phosphonopyruvate decarboxylases (25). The Km of sulfopyruvate was determined as 0.64 mM; the Vmax was 52 U/mg of protein. Phosphonopyruvate and pyruvate are not alternative substrates, despite the close similarity of the related enzymes. Phosphonopyruvate decarboxylase (PPD) [Scheme 1, Eq. (5)] is found in several Streptomyces species that form C–P compounds used as antibiotics or herbicides (fosfomycin, phosphinothricin, bialaphos). The initial step of C–P bond formation is catalyzed by phosphoenolpyruvate phosphonomutase, which creates phosphonopyruvate. The latter serves as substrate for PPD and undergoes decarboxylation to phosphonoacetaldehyde. Phosphonopyruvate decarboxylase enzyme activities have been determined in S. wedmorensis (26) and in S. viridochromogenes (27). The enzyme shows sequence similarity to various TDP enzymes that utilize pyruvate and related substrates. The use of other substrates than phosphonopyruvate or acyloin condensation activity has not been reported yet. Another highly interesting biotransformation is catalyzed by the bifunctional SHCHC synthase (SHCHC: 2-succinyl-6-hydroxy-2,4-cyclohexadiene1-carboxylic acid) and a-ketoglutarate decarboxylase [Scheme 1, Eq. (6)], readily enlarging the substrate range of TDP-dependent enzymes toward activated C–C double bonds (28–31). In mutant strains (sucA) of Bradyrhizobium japonicum that lack an active a-ketoglutarate dehydrogenase, another TDP-dependent activity that involves a-ketoglutarate but is coenzyme A independent was recently established (32). The product from a-ketoglutarate was shown to be succinic semialdehyde. Together with a succinate-semialdehyde dehydrogenase, this a-ketoglutarate decarboxylase [Scheme 1, Eq. (7)] may form an alternative pathway for a-ketoglutarate catabolism in this organism. A similar enzyme activity is found in the mitochondria of Euglena gracilis (33); a related bifunctional enzyme is part of the menaquinone biosynthetic pathway in bacteria (SHCHC; see earlier).

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Pyruvate oxidase from Escherichia coli (POX B) is a lipid-activated peripheral membrane flavoprotein enzyme that has a high degree of similarity to AHAS and glyoxylate carboligase. POX B decarboxylates pyruvate to acetate and to a lesser degree 2-oxobutanoate (‘‘a-ketobutyrate’’) to propionate (34). Pyruvate oxidase from Lactobacillus plantarum, however, differs from POX B because it is not activated by lipids. Furthermore, this enzyme produces from pyruvate acetyl phosphate and H2O2 instead, and 2-oxobutanoate is not a substrate for decarboxylation. Using knowledge based on the high-resolution crystal structure of the L. plantarum enzyme, Chang and Cronan altered the active site of the E. coli POX B and found mutant proteins that retained activity with 2-oxobutanoate but had nearly lost activity toward pyruvate (34). The authors therefore proposed the new name ‘‘a-ketobutyrate oxidase’’ (34) for the mutein. IV. C-1 METABOLISM Another very interesting aim is chain elongation through transformation of a C-1 unit (formaldehyde or equivalent), which is very difficult to perform selectively by chemical catalysis. Several TDP-dependent enzymes (TK, PDC, dihydroxyacetone synthase, glyoxylate carboligase) are known to catalyze such reactions, mostly in a nonasymmetric manner. Glyoxylate carboligase (GCL) from Escherichia coli catalyzes the condensation of two glyoxylate molecules to form tartronate semialdehyde plus CO2 [Scheme 2, Eq. (1)] (35). This reaction is mechanistically identical with acetolactate formation by AHAS. And GCL is also a flavoprotein, although FAD does not play a role in the catalysis (36). The TDP-bound intermediate is a hydroxymethyl group. However, GCL appears to be quite specific for glyoxylate because it does not cleave pyruvic acid (35,36). Dihydroxyacetone synthase is a transketolase-related enzyme found in methylotrophic yeasts such as Pichia and Hansenula polymorpha (37) or in methylotrophic bacteria (38). The enzyme transfers a C2 group from xylulose5-phosphate or, alternatively, from hydroxy pyruvate, after decarboxylation, onto formaldehyde [Scheme 2, Eq. (2)]. This reaction is also known from transketolases (2,4). Carbon dioxide fixation is a possible reaction for TDP-dependent enzymes catalyzing reversible reactions. This was recently shown for PDC [Scheme 2, Eq. (3)] (39). The a-oxidation pathway of 3-methyl-branched fatty acids, such as phytanic acids in mammals, involves a step in which the intermediate 2hydroxyphytanyl-CoA is cleaved into formyl-CoA and pristanal (40) (this volume, Chapter 29). This TDP-dependent step is catalyzed by the novel peroxisomal enzyme 2-hydroxyphytanoyl-CoA lyase [2-HPCL; Scheme 2,


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Scheme 2 Examples of the use of TDP-dependent enzymes in C-1 metabolism.

Eq. (4)] (40). The enzyme from Wistar rats was purified and the gene was isolated. Purified 2-HPCL had a specific activity of 558 mU/mg of protein with a Km of 15 lM for its substrate. 2-HPCL did not utilize the similar substrate 3-hydroxy-3-methylglutaryl-CoA from the mevalonate pathway. The native enzyme appeared to be a homotetramer with subunits of 63 kDa. The enzyme is homologous to bacterial oxalyl-CoA decarboxylases (e.g., from Oxalobacter formigenes) and to a similar enzyme from fungi and the nematode Caenorhabditis elegans. Oxalyl-CoA decarboxylases [OXC; Scheme 2, Eq. (5)] decarboxylate oxalyl-CoA to formyl-CoA and CO2 (41,42). These enzymes are found

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in bacteria and fungi and serve in a complete pathway for the degradation of oxalate, a toxic compound, especially for mammals. Oxalate-degrading bacteria as O. formigenes or Pseudomonas oxalaticus activate oxalate through reaction with succinyl-CoA, which delivers succinate and oxalyl-CoA-TDP. This enzyme-bound intermediate of OXC is then cleaved into carbon dioxide and formyl-CoA succinate complex. In the last step, succinyl-CoA is recycled and a formate is released (42). OXC from O. formigenes was purified and characterized (41) and its gene was cloned and analyzed (42). The substrate ranges of both 2-HPCL and OXC need to be characterized further; whether these enzymes can be utilized in acyloin condensation (with C1 compounds) also needs to be studied. V. KETONES AND IMINES AS ACCEPTORS From AHAS-catalyzed transformations it is known that ketones can serve as acceptor substrates [Scheme 3, Eq. (1)]. The recently described acetylacetoin synthase plays a role in the 2,3-butanediol cycle of several organisms (43), but the gene sequence is not available yet. The enzyme from Bacillus cereus is related to AHAS but differs because it transfers a C2 group stemming from diacetyl via a dihydroxyethyl-TPP intermediate onto a second molecule of diacetyl, thereby forming the branched product acetylacetoin [Scheme 3, Eq. (2)] (43,44). Whether the acetylacetoin synthase also accepts other ketones is not known yet. Liu and coworkers published the YerE-catalyzed ligation of activated acetaldehyde to a 4-keto-3,6-dideoxysugar, clearly emphasizing that ketones are promising acceptor substrates for TDP-dependent enzymes [Scheme 3, Eq. (3)] (45). Yersiniose A is a branched-chain sugar found in the O-antigen of Yersinia pseudotuberculosis VI and carries a two-carbon side chain that is derived from pyruvate. The gene product of yerE showed significant similarity (32% identical amino acid residues) to the large subunit of FADcontaining acetolactate synthases (45). Indeed, the purified recombinant YerE protein contained bound FAD. YerE has a calculated mass of 63,373 Da for each subunit and most likely exists as a homodimer. YerE transferred a C2 group from pyruvate onto an enzymatically prepared CDP precursor (a 3,6-dideoxy-4-hexulose) of yersiniose. The reaction product of YerE is then reduced in an NADPH-dependent step to yield CDP-yersiniose A. It is likely that other two-carbon branched sugars may derive from TDP-dependent reactions (45). In an ingenious follow-up study, Zhao and Liu (46) addressed the question of whether YerE tolerates a difluoro-group at the 4-keto position in order to study the mechanism of YerE. This attempt failed, however, due to a lack of substrate ambiguity of YerE (46).


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Scheme 3 Ketones and imines as acceptor substrates.

Frost and coworkers recently described the access to aminoDAHP via a transketolase-catalyzed formation of imino-erythrose-4-phosphate [Scheme 3, Eq. (4)]. This implies the use of a C–N bond in TDP-dependent catalysis (47). The application of imines as acceptor substrates in a benzoin-condensation-like reaction using thiazolium catalysts has long been known (48,49).


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As depicted in Scheme 3, Eq. (5), a ‘‘mixed’’ aminodeoxy benzoin can be obtained in high yield in a single step. Remarkably, in this case the corresponding benzoins are not observed and do not serve as substrates (50).

VI. MISCELLANEOUS REACTIONS Recently, Townsend and coworkers published an example of a TDP-dependent enzyme-catalyzed C–N bond formation [Scheme 4, Eq. (1)]. This enzyme from Streptomyces clavuligerus plays a key role in the formation of clavulanic acid, a h-lactamase inhibitor. The reaction is a C–N bond formation between D-glyceraldehyde-3-phosphate and L-arginine to give N2-(2-carboxyethyl)arginine (CEA) (51). The gene has been cloned (‘‘Orf2,’’ CEA-synthase) and found to encode a TDP-dependent enzyme (protein subunit size 60,907 Da) that shows about 29% sequence identity to acetolactate synthases from

Scheme 4 Miscellaneous TDP-dependent enzyme-catalyzed reactions.


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various sources. As reaction mechanism, the authors propose that D-glyceraldehyde-3-phosphate undergoes addition by the thiamine diphosphate ylide to form an intermediate that corresponds to a bound glycerol-3-phosphate. After h-elimination of hydroxide and ketonization, elimination of the phosphate group and addition of L-arginine may follow. Finally, addition of water would lead to the release of the product, CEA, and regeneration of the enzyme (51). Phosphoketolase, which is a rare TDP enzyme found mainly in some lactic acid bacteria, and Bifidobacteria catalyzes an irreversible TDP-dependent phosphorolytic reaction, e.g., cleaving fructose 6-phosphate plus inorganic phosphate to yield erythrose 4-phosphate and acetyl phosphate [Scheme 4, Eq. (2)] (52). Other putative acceptor substrates known from chemical benzoin- and acyloin-like condensations are Michael acceptors (Stetter reaction). An enzyme-catalyzed version of this reaction has been described in the literature, although the cited bakers’ yeast (BY) whole-cell biotransformation has not been elucidated in detail [Scheme 4, Eq. (3)]. It is assumed, that the depicted C–C bond formation between trifluoroacetaldehyde and the respective Michael acceptor is catalyzed by PDC (53).

VII. CONCLUSIONS AND OUTLOOK Most of the new enzymes and enzyme activities described in this chapter have been recognized due to investigations aiming to understand their physiological function in the respective organism. We propose that a detailed analysis of various TDP-dependent enzymes, with the focus on new transformations, which a priori do not have to be related to the respective physiological substrates or products, will open new perspectives in catalytic asymmetric synthesis. This assumption is nicely exemplified by the recent result obtained by Chipman, Barak, and coworkers that AHAS catalyzes the formation of (R)PAC even more efficiently than PDC (13). In the end, it should be possible to find an efficient matching TDP-dependent enzyme for the desired addition of various acyl anions to defined yet diverse electrophiles.

ACKNOWLEDGMENTS The work of the authors on TDP enzymes is supported by grants from the Deutsche Forschungsgemeinschaft through Sonderforschungsbereich 380 TP B21 (G.A.S.) and TP B27 (M.M.). The continuous support by H. Sahm and C. Wandrey is gratefully acknowledged.


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7 Ligand-Induced Conformational Changes in Thiamine Diphosphate–Dependent Enzymes: Comparison Between Crystal and Solution Structures Stephan Ko¨nig, Michael Spinka, Erik Fiedler, Georg Wille, and Johanna Brauer Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Michel H. J. Koch European Molecular Biology Laboratory, Hamburg Outstation, Germany Dmitri I. Svergun European Molecular Biology Laboratory, Hamburg Outstation, Germany, and Russian Academy of Sciences, Moscow, Russia

I. INTRODUCTION Reactions catalyzed by thiamine diphosphate (TDP) are widespread in all branches of metabolism, hence, not astonishingly, TDP-dependent enzymes are found in three of the four major enzyme classes (for a summary see Table 1). For the studies described here, we have chosen pyruvate oxidase from Lactobacillus plantarum (LpPOX, an oxidoreductase), transketolase from Saccharomyces cerevisiae (ScTK, a transferase), and pyruvate decarboxylase 93

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Table 1 Summary of Thiamine Diphosphate–Dependent Enzymes Oxidoreductases Phenylglyoxylate dehydrogenase (acylating) Pyruvate dehydrogenase (cytochrome) Pyruvate dehydrogenase (lipoamide) Pyruvate oxidase Oxoglutarate dehydrogenase (lipoamide) 3-Methyl-2-oxobutanoate dehydrogenase (lipoamide)

EC EC EC EC EC EC

Transferases Transketolase Formaldehyde transketolase Acetoin—ribose-5-phosphate transaldolase

EC 2.2.1.1 EC 2.2.1.3 EC 2.2.1.4

Lyases Pyruvate decarboxylase Benzoylformate decarboxylase Oxalyl-CoA decarboxylase Tartronate-semialdehyde synthase 2-Oxoglutarate decarboxylase Indolpyruvate decarboxylase 5-Guanidino-2-oxopentanoate decarboxylase Phosphoketolase Fructose-6-phosphate phosphoketolase Benzoin aldolase 2-Hydroxy-3-oxoadipate synthase Acetolactate synthase 1-Deoxy-D-xylulose 5-phosphate synthase Sulfoacetaldehyde lyase

EC EC EC EC EC EC EC EC EC EC EC EC EC EC

1.2.1.58 1.2.2.2 1.2.4.1 1.2.3.3 1.2.4.2 1.2.4.4

4.1.1.1 4.1.1.7 4.1.1.8 4.1.1.47 4.1.1.71 4.1.1.74 4.1.1.75 4.1.2.9 4.1.2.22 4.1.2.38 4.1.3.15 4.1.3.18 4.1.3.37 4.4.1.12

from brewer’s yeast (ByPDC, a lyase). LpPOX plays an important role in the energy metabolism of this organism by producing acetyl phosphate via oxidative decarboxylation of pyruvate in the presence of inorganic phosphate, oxygen, and the second cofactor, flavine adenine dinucleotide. ScTK is part of the pentose phosphate pathway and interconverts sugar monophosphates by ketol transfer. ByPDC is one of the enzymes involved in the alcoholic fermentation. It catalyzes the nonoxidative decarboxylation of pyruvate, yielding acetaldehyde and carbon dioxide. Substrate binding, the release of the first product, and the formation of the first intermediate, the socalled a-carbanion/enamine, is unique for all TDP-dependent enzymes, and a detailed general catalytic cycle has been established (1). Crystallographic models are not an absolute requirement for obtaining low-resolution models of the structure of proteins in solution on the basis of

Crystal and Solution Structures of TDP-Dependent Enzymes

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small-angle X-ray scattering patterns. However, when atomic resolution models are available, differences between the average structure in the crystal and in solution can easily be detected. For POX the two crystal structures available in the protein data bank (1POX, 1 POW) are those of the native holo enzyme, including all cofactors, but without ligands like substrates, activators, or inhibitors. The structure may be described as a dimer of dimers. The enzyme possesses a compact quaternary structure with extended interfaces between the subunits (2,3) (Fig. 1). In the case of ScTK, seven structures are deposited: the recombinant wild type (1TRK) (4), the variant H263A (1AYO) (5), complexes of ScTK with cofactor derivatives (1TKA, 1TKB, 1TKC) (6), with the central reaction intermediate, the a-carbanion of 2-(1,2-dihydroxy ethyl) TDP (1GPU) (7), and with an acceptor substrate (1NGS) (8). Figure 2 illustrates different views of the crystallographic model of holo ScTK (1TRK). This enzyme is a homodimer, with the cofactors TDP and metal ions bound at the interface between the subunits. Four crystal structures are available for the enzyme pyruvate decarboxylase, three from yeast [brewer’s yeast and Saccharomyces cerevisiae, 1PVD (9), 1PYD (10), 1QBP (11)] and one from the bacterium Zymomonas mobilis (1ZPD) (12). In all these cases the tetrameric structure is built up as a dimer of dimers, with the cofactors TDP and metal ions bound at the interface between subunits of each dimer (Fig. 3). All PDCs studied so far, except that from Zymomonas mobilis (ZmPDC), exhibit an exceptional kinetic property known as substrate activation (13–19). This is a rather slow process (for details see Ref. 13) that should be accompanied or caused by conformational changes of the protein (20). We suggest that the crystal structure of the complex of ByPDC with the substrate surrogate pyruvamide represents an activated enzyme form (1QBP) (11) (Fig. 3) and that the crystal structure of ByPDC in the absence of substrates or activators represents the native or inactive enzyme form. The models of ScTK, LpPOX, and ByPDC, illustrated in Figures 1–3, have been compared in detail elsewhere (21) and are used as a starting point in the refinement procedure described here. Small-angle X-ray solution scattering (SAXS) data for all enzyme solutions were collected via the X33 camera of the European Molecular Laboratory in HASYLAB on the storage ring DORIS of the Deutsches Elektronensynchrotron at Hamburg. Measurements were done at camera lengths of 2.9 and 3.4 m, covering a range of momentum transfer s of 0.1–2.3 nm1 (s= (4k sin u)/E, 2u is the scattering angle, and E= 0.15 nm, the wavelength of the X-rays). Protein concentrations were 4–5 mg/mL. The buffer conditions guaranteed the pH optimum for catalytic activity and avoided pH shifts after addition of ligands. Initial treatment of the scattering data was performed with the programs Sapoko (DI Svergun, MHJ Koch, unpublished data) and Gnom (22) (for details, see http://www.embl-ham-

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Figure 1 Crystallographic model of a stablized mutant of the recombinant wild type of pyruvate oxidase from Lactobacillus plantarum (1POX). The structure is shown as a solid ribbon, with different shading of the subunits; cofactors are not included; middle cell, 90j rotation of the upper structure around the y-axis; lower cell, 90j rotation of the middle structure around the x-axis. (From Ref. 3.)


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Figure 2 Crystallographic model of the recombinant wild type of transketolase from Saccharomyces cerevisiae (1TRK). Same mode of presentation as in Figure 1. (From Ref. 4.)

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Figure 3 Crystallographic models of different pyruvate decarboxylases. Left column, pyruvate decarboxylase from brewer’s yeast (1PVD, from Ref. 9), middle column, complex of pyruvate decarboxylase from brewer’s yeast strain and the artificial activator pyruvamide (1QBP, from Ref. 11); right column, recombinant wild type of pyruvate decarboxylase from Zymomonas mobilis (1ZPD, from Ref. 12). Same mode of presentation as in Figure 1.


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burg.de/ExternalInfo/Research/Sax/). The rigid-body refinement was performed using the program Massha (23). Scattering patterns were calculated from the crystallographic models using the program Crysol (24), which surrounds the enzyme macromolecule with a 0.3-nm-thick hydration layer with an adjustable density. For the rigid-body refinement the crystallographic structure models were oriented in such way that the twofold axis transforming one half-molecule into the other (these are monomers in the case of ScTK and dimers in the case of ByPDC and LpPOX) coincided with the y-axis. The refined parameters were the rotation angles of one half-molecule around the center of mass and its shift along the z-axis. An automated fit procedure was performed to find the values of these parameters providing the best fit to the experimental scattering data (indicated as discrepancy m; for a detailed description of the fitting procedure, see Refs. 25 and 26). Scattering patterns were recorded for all enzymes in the absence and the presence of saturating concentrations of ligands. In this study the following ligands were used: for LpPOX the substrate pyruvate, for ScTK the artificial donor substrate 3hydroxypyruvate and the synthetic intermediate D,L-2-(1,2-dihydroxy ethyl)TDP (DHETDP), respectively, and for ByPDC the substrate surrogate and artificial activator pyruvamide. The chosen experimental conditions prevented significant consumption of the ligands during measurements by the catalytically active enzymes: the limited amount of oxygen in case of LpPOX, the lack of an acceptor substrate and the relatively high stability of DHETDP in case of ScTK, and the decarboxylase-resistant amide bond of pyruvamide in the case of ByPDC.

II. RESULTS A. Comparison of the Crystal Structure and Solution Structure and the Effect of the Substrate Pyruvate on the Solution Structure of Pyruvate Oxidase from Lactobacillus plantarum (LpPOX) Comparison of the experimental data of LpPOX processed by Gnom with the curve calculated from the crystallographic model revealed significant differences in the s range of 0.8–1.2 nm1, pointing to differences between the quaternary structure of LpPOX in the crystal and in solution (Fig. 4). The fit was improved by rigid-body refinement, and the m-value dropped by about 0.17. This corresponded to a shift of 4–5 A˚ and a rotation of 11–14j of one dimer relative to the other, indicating that the quaternary structure of LpPOX in solution is less compact than in the crystal (Fig. 5). The same result was obtained by comparing crystal and solution structures when the scattering patterns had been recorded under crystallizaton-like

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Figure 4 Experimental small-angle X-ray solution scattering pattern and fits of the curves of the original crystallographic model and of the solution structure model obtained by rigid-body refinement for pyruvate oxidase from Lactobacillus plantarum (A) in the absence and (B) the presence of pyruvate. Open circles, experimental data; dark grey solid line, processed curve by the program Gnom; black dashed line, scattering curve calculated from the crystallographic model using the program Crysol; black solid line, scattering curve calculated from the refined solution structure model using the program Massha.


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Figure 5 Comparison of the crystallographic model of the recombinant wild type of pyruvate oxidase from Lactobacillus plantarum (1POX, left column) and the corresponding solution structure models obtained by rigid-body refinement using the automated fit procedure of the program Massha in the absence (middle column) and presence (right column) of the substrate pyruvate. Same mode of presentation as in Figure 1.

conditions (25). Addition of the substrate pyruvate to solutions of LpPOX did not change the calculated solution structure significantly. The same extent of translation (4–5 A˚) and rotation (11–14j) of one dimer relative to the other in comparison to the crystal structure of the enzyme is necessary to obtain the best fit to the experimental scattering data (Dm= 0.13–0.19, Figs. 4B and 5). This means, that the presence of the natural substrate pyruvate did

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not induce any changes detectable by our methods in the solution structure of LpPOX. B. Comparison of the Crystal Structure and Solution Structure and the Effect of the Artificial Donor Substrate 3-Hydroxy Pyruvate and the Chemically Synthesized Reaction Intermediate D,L-2-(1,2-Dihydroxy Ethyl)-TDP (DHETDP) on the Solution Structure of Transketolase from Saccharomyces cerevisiae (ScTK) The processed experimental data and the curve calculated from the crystallographic model were also not identical in the case of unliganded ScTK, as illustrated in Figure 6A. Deviations between the plots calculated with Gnom and Crysol occurred predominately in the s region of 11.5 nm1. In the resulting solution structure model of unliganded ScTK, the distance between the centers of mass is increased by 5–5.5 A˚ and the monomers were only slightly rotated by about 2–4j. The quaternary structure of ScTK in solution is less compact than in the crystal (Fig. 7). Addition of the artificial donor substrate 3-hydroxy pyruvate increased the difference between the experimental scattering data and the curve calculated from the crystallographic model (Fig. 6B). Consequently, the decrease in the discrepancy m during the rigid-body refinement was greater in the liganded (0.1–0.2) than in the unliganded state (0.05–0.1) of ScTK. The corresponding solution structures exhibited a similar shift of 4–5 A˚, but the rotation angle was increased by 5–8j (Fig. 7). The differences between crystal and solution models were even more pronounced upon addition of the reaction intermediate DHETDP, and the discrepancies for the fit of the curve calculated from the crystallographic model and the experimental data were rather high. Refinement reduced these values by about 0.4, but significant deviations remained especially at higher svalues (Fig. 6C). The models of the solution structure of the DHETDP-ScTK

Figure 6 Experimental small-angle X-ray solution scattering pattern and fits of the curves of the original crystallographic model and of the solution structure model obtained by rigid-body refinement for transketolase from Saccharomyces cerevisiae without any ligand (A), in the presence of 3-hydroxy pyruvate (B), and in the presence of D,L-2-(1,2-dihydroxy ethyl)-TDP (C). Open circles, experimental data; dark grey solid line, processed curve by the program Gnom; black dashed line, scattering curve calculated from the crystal structure model using the program Crysol; black solid line, scattering curve calculated from the refined solution structure model using the program Massha.


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Figure 7 Comparison of the crystallographic model of the recombinant wild type of transketolase from Saccharomyces cerevisiae (1TRK; left column) and the corresponding solution structure models obtained by rigid-body refinement using the automated fit procedure of the program Massha in the absence (second column) of any ligand, in the presence of 3-hydroxy pyruvate (third column), and in the presence of D,L-2-(1,2-dihydroxy ethyl)-TDP (right column). Same mode of presentation as in Figure 1.

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complex displayed the same shift as in the unliganded and the substrateliganded state (4–5 A˚), but the rotation angle between the monomers increased to 12–13j (Fig. 7). C. Comparison of the Crystal Structure and Solution Structure and the Effect of the Artificial Activator Pyruvamide on the Solution Structure of Pyruvate Decarboxylase from Brewer’s Yeast (ByPDC) The largest deviation between experimental scattering data and curves calculated from the crystallographic model of unliganded enzyme states was found for the enzyme ByPDC (Fig. 8A). The best fit of the refined model decreased the discrepancy m by about 0.3. Surprisingly, the solution structure is more compact than the crystal structure, visualized by a shift of one dimer toward the other of 5–6.5 A˚. An additional rotation of one dimer relative to the other of 9–12j resulted in a structure illustrated in Figure 9. Addition of pyruvamide to solutions of the holo-ByPDC changed the scattering pattern significantly, as shown previously (27) and in Figure 8B. There was a better fit of the experimental data and the calculated curve derived from the crystal structure at lower s-values (0.7–1.1 nm1), but deviations increased around 1.5 nm1 (Fig. 8B). Rigid-body refinement decreased the discrepancy by 0.1 only, and the distance between the centers of mass of dimers of the crystal and solution structure remained unchanged. The refinement resulted in a large rotation by 40–70j of one dimer relative to the other. The dimer orientation in the solution structure of the pyruvamide-activated enzyme resembles more closely the crystal structures of LpPOX and ZmPDC than that of ByPDC (compare Figs. 1–3 with Fig. 9). The capability of the refinement program package Massha was clearly demonstrated by the fact that it was also possible to use the crystal structures of pyruvamide-activated ByPDC (11,28) and of ZmPDC (12) (Fig. 3) as initial models. Although the discrepancies between the fitted experimental data and the calculated curves of the crystal structures using the program Crysol were rather high, the m-values dropped down to the same values as in the case of native ByPDC or tended to be even lower after running the Massha program (Fig. 8C). Furthermore, the values for the translation of dimers within the tetramers were exactly the same in all three cases, namely, that of the crystal structure model of ByPDC, and the relative rotation of dimers was in the same range (35–80j). III. DISCUSSION Significant differences were found between the crystal structure and solution structure of all TDP-dependent enzymes studied here. Translations of half-

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molecules by 4–6 A˚ and corresponding rotations of 3–12j gave the best fit to the experimental solution scattering curves. The rotation of the monomer in ScTK is significantly smaller than that observed for the two other enzymes, and the distance between dimers in ByPDC is reduced by 4–6 A˚. The structural differences between solution and crystal states may thus be related to the compactness of the quaternary structure of the enzymes; the larger the subunit interface area, the smaller the differences between crystal and solution structures. Enzymes with rather compact quaternary crystal structures, like LpPOX (2,3) and ScTK (4,29), appear expanded in solution; vice versa, those with a very small interface between subunits (especially dimers), like ByPDC, are compressed in solution. An earlier comparison between the crystallographic and solution structures of ByPDC and ZmPDC yielded similar results (25). Hydration can in principle be excluded as a general reason for this phenomenon, because the program Crysol adds a hydration layer to the crystallographic model. Furthermore, the scattering patterns calculated in this way fit the corresponding experimental data very well at small angles. This demonstrates that the quaternary structure of these oligomeric enzymes is the same in the crystal and solution states (25). Figures 4, 6, and 8 illustrate that significant deviations between fits of experimental data and calculated curves occur at higher scattering angles, a region sensitive to intramolecular interactions, for instance, caused by changes of the relative arrangement of the subunits. The Massha program enables one to analyze possible subunit rearrangements on the basis of experimental scattering data using rigid-body refinement and was applied in this study to investigate ligand-induced conformational changes. In contrast with the general changes between crystal and solution states of the various TDP-dependent enzymes, there is no common effect of enzymespecific ligands on the enzyme conformation. Addition of the natural sub-

Figure 8 Experimental small-angle X-ray solution scattering pattern and fits of the curves of the original crystallographic model and of the solution structure model obtained by rigid-body refinement for pyruvate decarboxylase from brewer’s yeast in the absence (A) and in the presence of the artificial activator pyruvamide (B). Open circles, experimental data; dark grey solid line, curve processed by the program Gnom; black dashed line, scattering curve calculated from the crystal structure model using the program Crysol; black solid line, scattering curve calculated from the refined solution structure model using the program Massha. (C) Comparison of calculated curves of solution structures after the refinement of different crystallographic models. Grey line, ByPDC; black dashed line, pyruvamide activated ByPDC; black solid line, ZmPDC.

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Figure 9 Comparison of the crystallographic model of pyruvate decarboxylase from brewer’s yeast (1PVD, left column) and the corresponding solution structure models obtained by rigid-body refinement using the automated fit procedure of the program Massha in the absence (middle column) and in the presence of the artificial activator pyruvamide (right column). Same mode of presentation as in Figure 1.

strate pyruvate to LpPOX does not alter the dimer arrangement significantly as compared to the unliganded state. The artificial donor substrate 3hydroxypyruvate and the intermediate-like compound 2-(1,2-dihydroxy ethyl)-TDP increase the rotation angle of the subunits of ScTK from 3j to 8j and 13j, respectively. Dramatic effects on the solution structure of ByPDC result from the addition of the artificial activator and substrate surrogate


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pyruvamide. The compression of the unliganded state is neutralized, and a large relative rotation of the dimers of the enzyme is required to fit the data. All ligands used are very specific for the individual enzymes, and no nonspecific effects during the binding should be expected. Only very small changes, if any, are observed for LpPOX and ScTK, and specific effects of substrate binding have not been described so far with other experimental methods. Cocrystallization or crystal soaking of ScTK with various substrates and cofactor derivatives, respectively, does not change the overall crystal structure of the enzyme (6–8,30). In contrast, a number of kinetic studies on the substrate activation behavior of ByPDC (13– 19) suggest extensive conformational changes of the enzyme during this process. First hints of distinct inactive and substrate-activated states came from cross-linking and small-angle X-ray solution scattering experiments (20,27). Crystal structure analysis of the native and pyruvamide-activated states of ByPDC confirmed these results (11,28) (Fig. 3). The main differences between both crystal structures are the rearrangements of the dimers within the tetramer, resulting in a 30j rotation and the fixing of two loop regions near the active sites of the enzyme. Our current results point in the same direction—the rearrangement of the dimers by an even more pronounced rotation by 40–70j. Whether the ZmPDC structure with 80j rotation angle between dimers resembles the fully activated ByPDC structure is still not clear, and the question it was not expected to be answered here. But the obtained solution structure of pyruvamide-activated ByPDC supports our hypothesis that dimer reorientation is the structural prerequisite and/or result of the substrate activation. The results demonstrate that the application of the procedure of rigidbody refinement, together with small-angle X-ray solution scattering data, is a promising way to obtain information on protein structures in solution when crystal structures are available. The lower discrepancies (given as Dm in Table 2) for all solution structure models obtained by rigid-body refinement illustrate that these models represent the native solution structure of all studied enzyme forms better than do the crystallographic models. The limitations of the method should, however, be kept in mind when interpreting the resulting models. Information on intramolecular interactions can only be drawn from the higher s range of scattering patterns (Figs. 4, 6, and 8). Unfortunately, in this range the accuracy of the experimental data is lower than at smaller angles. Careful data collection and interpretation are therefore decisive for the applicability of the method. Measurements at various camera lengths and high protein concentrations can improve the reliability of the scattering pattern at high angles, depending on the molecular mass of the protein, but possible effects of intermolecular interactions should be taken into account (31,32). Even the best fit of the calculated solution models still

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Table 2 Parameters of Solution Structure Models of the Enzymes LpPOX, ScTK, and ByPDC in the Absence and Presence of Ligands Obtained by Rigid-Body Refinement of the Corresponding Crystallographic Structures Using the Automated Fit Procedure of the Program Massha ScTK LpPOX

Dm Translation (A˚) Rotation. (j) r.m.s. (A˚)

ByPDC

No ligand

Pyruvate

No ligand

Hydroxy pyruvate

0.15–0.19 4–5

0.13–0.19 4–5

0.05–0.1 5–5.5

0.1–0.2 4–5

0.4 4–5

0.3 (5–6.5)

0.1 0

11–14 7.0–7.5

11–14 6.5–7.2

2–4 2.6–2.8

5–8 2.3–4.6

12–13 3.6–5.1

9–12 5.4–6.2

40–70 19.3

DHETDP

No ligand

Pyruvamide

The values are means of at least five indiviual experiments. Dm= mcrysol– mmassha; translation, shift along the z–axis of the half–molecule; rotation, around the center of mass of one half-molecule; r.m.s., root mean square displacement of the atomic coordinates of the crystal and solution structure model.

deviates from the experimental data, especially in the case of ScTK. This is caused by the breakdown of the rigid-body assumption for the half-molecule corresponding to one subunit in the case of ScTK and the dimers in the case of LpPOX and ByPDC, respectively. Obviously, in solution the outer parts of a macromolecule are more flexible then the core, regardless of the subunit they belong to. The mobility of individual domains within one subunit may also vary. This flexibility is not taken into account during the rigid-body refinement procedure and thus is also not represented in the resulting solution structure models. The apparent atomic resolution of these models should not create the illusion that they represent more than a better approximation to the solution structure obtained by an effectively low-resolution refinement in terms of rotation and translation of half-molecules along one axis. The models should thus not be used in detailed comparison with the original crystallographic model, but rather as a measure of the differences in the conformational spaces explored by the macromolecules in the crystal and in solution. REFERENCES 1. 2. 3.

A Schellenberger, G Hu¨bner, H Neef. Cofactor designing in functional analysis of thiamin diphosphate enzymes. Meth Enzymol 279:131–146, 1997. YA Muller, GE Schulz. Structure of the thiamine-and flavin-dependent enzyme pyruvate oxidase. Science 259:965–967, 1993. YA Muller, G Schumacher, R Rudolph, GE Schulz. The refined structures of a


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stabilized mutant and of wild-type pyruvate oxidase from Lactobacillus plantarum. J Mol Biol 237:315–335, 1994. M Nikkola, Y Lindqvist, G Schneider. Refined structure of transketolase from Saccharomyces cerevisiae at 2.0-A˚ resolution. J Mol Biol 238:387–404, 1994. C Wikner, U Nilsson, L Meshlkina, C Udekwu, Y Lindqvist, G Schneider. Identification of catalytically important residues in yeast transketolase. Biochemistry 36:15643–15649, 1997. S Ko¨nig, A Schellenberger, H Neef, G Schneider. Specifity of coenzyme binding in thiamin diphosphate-dependent enzymes. Crystal structures of yeast transketolase in complex with analogs of thiamin diphosphate. J Biol Chem 269: 10879–10882, 1994. E Fiedler, S Thorell, T Sandalova, R Golbik, S Ko¨nig, G Schneider. Snapshot of a key intermediate in enzymatic thiamin catalysis: crystal structure of the acarbanion of (a,h-dihydroxyethyl)-thiamin diphosphate in the active site of transketolase from Saccharomyces cerevisiae. Proc Natl Acad Sci USA 99:591– 595, 2002. U Nilsson, L Meshalkina, Y Lindqvist, G Schneider. Examination of substrate binding in thiamin diphosphate–dependent transketolase by protein crystallography and site-directed mutagenesis. J Biol Chem 272:1864–1869, 1997. P Arjunan, T Umland, F Dyda, S Swaminathan, W Furey, M Sax, B Farrenkopf, Y Gao, D Zhang, F Jordan. Crystal structure of the thiamin diphosphate– dependent enzyme pyruvate decarboxylase from the yeast Saccharomyces cerevisiae at 2.3-A˚ resolution. J Mol Biol 256:590–600, 1996. F Dyda, W Furey, S Swaminathan, M Sax, B Farrenkopf, F Jordan. Catalytic centers in the thiamin diphosphate–dependent enzyme pyryvate decarboxylase at 2.4-A˚ resolution. Biochemistry 32:6165–6170, 1993. G Lu, D Dobritzsch, S Baumann, G Schneider, S Ko¨nig. The structural basis of substrate activation in yeast pyruvate decarboxylase—a crystallographic and kinetic study. Eur J Biochem 267:861–868, 2000. D Dobritzsch, S Ko¨nig, G Schneider, G Lu. High-resolution crystal structure of pyruvate decarboxylase from Zymomonas mobilis. Implications for substrate activation in pyruvate decarboxylases. J Biol Chem 273:20196–20204, 1998. G Hu¨bner, R Weidhase, A Schellenberger. The mechanism of substrate activation of pyruvate decarboxylase: a first approach. Eur J Biochem 92:175–181, 1978. G Gish, T Smyth, R Kluger. Thiamin diphosphate catalysis. Mechanistic divergence as a probe of substrate activation of pyruvate decarboxylase. J Am Chem Soc 110:6230–6234, 1988. A Schellenberger, G Hu¨bner, S Ko¨nig, S Flatau, H Neef. Substrate activation of pyruvate decarboxylase—mechanistic aspects. Nova Acta Leopold 61:225–242, 1989. I Baburina, Y Gao, Z Hu, F Jordan, S Hohmann, W Furey. Substrate activation of brewer’s yeast pyruvate decarboxylase is abolished by mutation of cysteine 221to serine. Biochemistry 33:5630–5635, 1994. A Dietrich, S Ko¨nig. Substrate activation behavior of pyruvate decarboxylase from Pisum sativum cv. Miko. FEBS Lett 400:42–44, 1997.

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18. J Wang, R Golbik, B Seliger, M Spinka, K Tittmann, G Hu¨bner, F Jordan. Consequences of a modified putative substrate-activation site on catalysis by yeast pyruvate decarboxylase. Biochemistry 40:1755–1763, 2001. 19. F Krieger, M Spinka, R Golbik, G Hu¨bner, S Ko¨nig. Pyruvate decarboxylase from Kluyveromyces lactis—an enzyme with an extraordinary substrate activation behavior. Eur J Biochem 269:3256–3263, 2002. 20. S Ko¨nig, G Hu¨bner, A Schellenberger. Cross-linking of pyruvate decarboxylase. Characterization of the native and substrate-activated states. Biomed Biochim Acta 49:465–471, 1990. 21. YA Muller, Y Lindqvist, W Furey, GE Schulz, F Jordan, G Schneider. A thiamin diphosphate binding fold revealed by comparison of the crystal structures of transketolase, pyruvate oxidase and pyruvate decarboxylase. Structure 1:95–103, 1993. 22. DI Svergun. Determination of the regularization parameter in indirect transform methods using perceptual criteria. J Appl Crystallogr 25:495–503, 1992. 23. PV Konarev, MV Petoukhov, DI Svergun. MASSHA—a graphics system for rigid-body modelling of macromolecular complexes against solution scattering data. J Appl Cryst 34:527–532, 2001. 24. DI Svergun, C Barbareto, MHJ Koch. Crysol—a program to evaluate X-ray solution scattering of biological macromolecules from atomic coordinates. J Appl Cryst 28:768–773, 1995. 25. DI Svergun, MV Petoukhov, MHJ Koch, S Ko¨nig. Crystal versus solution structures of thiamine diphosphate–dependent enzymes. J Biol Chem 275:297– 302, 2000. 26. S Ko¨nig, DI Svergun, VV Volkov, LA Feigin, MHJ Koch. Small-angle X-ray scattering studies on ligand-induced subunit interactions of the thiamine diphosphate-dependent enzyme pyruvate decarboxylase from different organisms. Biochemistry 37:5329–5334, 1998. 27. G Hu¨bner, S Ko¨nig, A Schellenberger, MHJ Koch. An X-ray solution scattering study of the cofactor and activator-induced structural changes in yeast pyruvate decarboxylase (PDC). FEBS Lett 266:17–20, 1990. 28. G Lu, D Dobritzsch, S Ko¨nig, G Schneider. Novel tetramer assembly of pyruvate decarboxylase from brewer’s yeast observed in a new crystal form. FEBS Lett 403:249–253, 1997. 29. Y Lindqvist, G Schneider, U Ermler, M Sundstro¨m. Three-dimensional structure of transketolase, a thiamine diphosphate–dependent enzyme, at 2.5-A˚ resolution. EMBO J 11:2373–2379, 1992. 30. U Nilsson, Y Lindqvist, R Kluger, G Schneider. Crystal structure of transketolase in complex with thiamine thiazolone diphosphate, an analogue of the reaction intermediate, at 2.3-A˚ resolution. FEBS Lett 326:145–148, 1993. 31. S Ko¨nig, MHJ Koch. The effect of high protein concentrations on the pHdependent scattering behavior of the enzyme pyruvate decarboxylase from brewer’s yeast. Hasylab Ann Report, 909–910, 1998. 32. S Ko¨nig, MHJ Koch. The effect of high protein concentrations on the SAX scattering behavior of two different species of pyruvate decarboxylase. Hasylab Ann Report, 991–992, 1999.

8 Enantioselective Syntheses of Hydroxy Ketones via Benzoylformate Decarboxylaseand Benzaldehyde LyaseCatalyzed C–C Bond Formation Bettina Lingen and Martina Pohl Heinrich-Heine-Universita¨t Du¨sseldorf, Ju¨lich, Germany Ayhan S. Demir Middle East Technical University, Ankara, Turkey Andreas Liese and Michael Mu¨ller Forschungszentrum Ju¨lich GmbH (Research Centre Ju¨lich), Ju¨lich, Germany

I. INTRODUCTION Various TDP-dependent a-keto acid decarboxylases have been described as catalyzing C–C bond formation and/or cleavage (1). Here we want to draw attention to some concepts based on the investigation of reactions catalyzed by the enzymes benzoylformate decarboxylase (BFD) and benzaldehyde lyase (BAL), the genes of which were cloned and the proteins overexpressed in recombinant E. coli strains. An overview is given with respect to the development of muteins with improved properties, such as higher carboligase 113

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activity in the presence of water-miscible organic solvents and altered substrate range, that were created by rational protein design as well as by directed evolution. II. BENZOYLFORMATE DECARBOXYLASE: WILD-TYPE ENZYMES AND MUTEINS The potential of benzoylformate decarboxylase (BFD, E.C. 4.1.1.7) to catalyze C–C bond formation was first reported by Wilcocks et al. using crude extracts of Pseudomonas putida (2). They observed the formation of (S)2-hydroxy-1-phenyl-propanone [(S)-2-HPP] when benzoylformate was decarboxylated in the presence of acetaldehyde. Advantageously, aldehydes without a previous decarboxylation step can be used instead of the corresponding, more expensive a-keto acids (3). As depicted in Table 1, BFD is able to bind a broad range of different aromatic and heteroaromatic aldehydes to TDP prior to ligation to acetaldehyde (4). Best results with respect to the enantiomeric excess (ee) of the resulting 2-hydroxy ketones were obtained with metasubstituted benzaldehydes. Using these substrates, the ee increased to more than 99%, indicating that the sterical demand and electronic properties of the substituent play a decisive role in both conversion rate and enantiomeric excess (ee). ortho-Substituted benzaldehyde derivatives, except 2-fluoroben-

Table 1 Wild-Type BFD-Catalyzed Carboligation Toward (S)-2-HPP Derivatives

Ar Ph 3-MeOC6H4 3-iPrOC6H4 3,5-di-MeOC6H3 2-Naphthyl 3-Pyridinyl 2-(5-Methyl)furanyl 2-Thiophenyl a

Yield (%)

eeb (%)

Configuration

90 97 91 40 32 65a 32a 50a

92 96 >99 97 88 87 86 83

(S) (S) (S) (S) (S) (S) (S) (S)

Conversion (%), determined by GCMS and HPLC. ee = enantiomeric excess. Source: Refs. 4 and 5.

b

Enantioselective Syntheses of Hydroxy Ketones

115

zaldehyde, are only poorly accepted as donor substrates by the wild-type enzyme, probably due to sterical hindrance (5). Being aware that meta- and para-substituted aromatic aldehydes provide the highest ee values with good to excellent conversion rates, we successfully subjected dialdehydes to the BFD-catalyzed coupling reaction, resulting in the stereospecific formation of diadducts (6). Reaction of isophthalaldehyde (1) and excess acetaldehyde in the presence of BFD gave diadducts (S,S)-2 (ee > 99%) and meso-2 in a ratio of 94:6 and a combined yield of 82% [Scheme 1, entry (1)]. In the case of terephthalaldehyde (3) as a substrate, product formation was considerably slower. Even after prolonged reaction time, no more than 14% of the diadduct (S,S)-4 (ee > 99%) and meso-4 were obtained in a combined yield after isolation [Scheme 1, entry (2)]. Not only aromatic aldehydes, but also cyclic aliphatic and conjugated olefinic aldehydes are accepted as donor substrates by BFD. Thus, for the first time we could demonstrate the BFD-mediated stereoselective cross-coupling

Scheme 1 BFD-catalyzed carboligation of isophthalaldehyde (1) [terephthalaldehyde (3)] and acetaldehyde, yielding (S,S)-2 [(S,S)-4]. (From Ref. 6.)

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Scheme 2 BFD-catalyzed carboligation of cyclohexenecarbaldehyde (5) and acetaldehyde. (From Ref. 5.)

of two different aliphatic substrates, cyclohexane carbaldehyde and acetaldehyde, or olefinic and aliphatic substrates (Scheme 2) (4). In contrast to the large variety of diverse aldehydes that can be used as donor substrates, wild-type BFD does not tolerate a modification of the methyl group of acetaldehyde in the case of aliphatic acceptor aldehydes. Apart from acetaldehyde, BFD shows activity with aromatic and heteroaromatic aldehydes as the acceptor substrate, forming enantiomerically pure (R)-benzoin and derivatives (Table 2) (7). A. Control of Enantioselectivity in C–C Bond Formations Catalyzed by Wild-Type BFD The enantioselectivity of asymmetric C–C bond formations catalyzed by wild-type BFD can be influenced by three different parameters: temperature, benzaldehyde concentration, and electronic and steric properties of aromatic

Table 2 Wild-Type BFD-Catalyzed Benzoin Condensation to the Corresponding (R)-Benzoins on a Preparative Scale

Ar

Yield (%)

eea (%)

Configuration

70 68 18 69 62 50

>99 >99 >99 >99 94 96

(R) (R) (R) (R) (R) (R)

Ph 2-FC6H4 3-MeOC6H4 4-MeC6H4 2-Furanyl 2-(5-Methyl)furanyl a

ee = enantiomeric excess. Source: Ref. 7.


117

substituents on the donor aldehyde. In the case of the C–C bond formation between benzaldehyde and acetaldehyde, kinetic investigations have shown that the enantioselectivity is increased by lowering the temperature. Surprisingly, the enantiomeric excess of the resulting hydroxyketone is also increased if the initial concentration of benzaldehyde is lowered (Fig. 1). The transition states of the catalytic reaction were modeled based on the X-ray crystallographic data published for wild-type BFD (8), with the benzaldehyde bound to the C-2 of the thiazolium in thiamine diphosphate (4). As a result it could be shown that two different orientations of the previously described intermediate are possible in the active site, leading to the opposite enantiomers. These two conformations are believed to be in a kinetic equilibrium that can be shifted by temperature and the benzaldehyde concentration, resulting in different values of enantiomeric excess for (S )-2-HPP. The third parameter that influences the enantiomeric excess is the electronic and steric properties of different substituents on the benzaldehyde. Different substituents in the para and meta-positions of benzaldehyde demonstrate a broad range in the resulting enantiomeric excess of the carboligation products. It was demonstrated that in the case of para-substituents, the enantiomeric excess follows the Hammett correlation (9). The Hammett equation correlates the ratio of the acid dissociation constants of benzoic acids with the rates of alkaline hydrolysis of benzoic esters. In other words, it denotes the electronic properties of the different substituents. In the case of

Figure 1 Enantiomeric excess as a function of benzaldehyde concentration and temperature (50 mM Kpi, pH 7.0, 500 mM acetaldehyde).

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the enantioselective carboligation catalyzed by wild-type BFD, the logarithm of the ratio of S- to R-enantiomer can be correlated with the j-constant of the Hammett equation (Fig. 2) (10). The term log (S/R) is proportional to DDG#: logðS=RÞ ¼ DDG# =RT

ð1Þ

DDG# ¼ ðDG#S DG#R Þ

ð2Þ

DG# ¼ k eE=RT

ð3Þ

with and

In the case of meta-substituted benzaldehydes, no significant correlation with the Hammett constant can be observed (Fig. 3). It appears that steric parameters dominate the electronic ones in this case. The increase of the enantiomeric excess with electron-donating substituents in the case of parasubstituted benzaldehydes can be explained by a stabilization of the edge-toface interactions of the phenyl residues of the phenylalanines 464 and 397 in the active site of BFD with the phenyl residue of the aldehyde donor. Knowledge of the dependence of the enantiomeric excess on substituent effects enables the synthetic chemist to select the synthetic route leading to the highest ee possible.

Figure 2 Hammett plot for para-substituted benzaldehydes.


119

Figure 3 Hammett plot for meta-substituted benzaldehydes.

B. Asymmetric Cross-Benzoin Condensation with BFD H281A Starting from the assumption that aldehydes not accepted as donor substrates might still be suitable acceptor substrates, and vice versa, we performed a mixed enzyme–substrate screening in order to identify a biocatalytic system for the asymmetric cross-carboligation of aromatic aldehydes. For this purpose, 2-chloro- (7a), 2-methoxy- (7b), and 2-methylbenzaldehyde (7c) were reacted with different enzymes in combination with benzaldehyde (Scheme 3) (11). The three ortho-substituted benzaldehyde derivatives 7a–c were chosen as putative selective acceptor substrates particularly because of their inability to form symmetrical benzoins through the wild-type-BFD– catalyzed reaction, meaning that these compounds are not accepted as donor substrates by this enzyme. The BFD-mutant BFD H281A (12), which was created by directed mutagenesis, was identified as a potent catalyst, resulting in the formation of the mixed benzoins 2V-methoxybenzoin (8b) and 2Vmethylbenzoin (8c), accompanied by (R)-benzoin as the major product. In the case of 2-chlorobenzaldehyde (7a) as acceptor substrate, the unsymmetrical benzoin (R)-8a (yield 74%, ee > 99%) represents the major product (11). As expected, the 2,2V-disubstituted benzoin 9 or the mixed benzoin 10 substituted in the 2-position was not generated in any of these reactions,

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Scheme 3 Asymmetric synthesis of mixed benzoins 8a-c by use of BFD H281A.

revealing that the ortho-substituted benzaldehydes 7a–c react selectively as acceptors. Subsequently, we extended our concept to selective donor molecules. With 2-chloro-benzaldehyde as the selective acceptor, a vast variety of unsymmetrical benzoins was accessible, among which 8d, 8e, and 8f (Structure 1) were obtained selectively, proving that 4-(trifluoro-methyl)-benzaldehyde, 4-bromobenzaldehyde, and 3-cyanobenzaldehyde were selective donor substrates for BFD H281A (11).

Although the racemic cross-benzoin condensation catalyzed by cyanide under forcing conditions has been known since the end of the 19th century (13), the possibility that the observed formation of specific mixed benzoins, at least in some cases, is a result of thermodynamic equilibration rather than selective formation cannot be excluded (14). The results of the asymmetric cross-benzoin condensation (11) prove beyond any doubt the selective formation of the mixed benzoins 8 due to the selective donor or selective acceptor behavior of the corresponding aromatic aldehydes. C. BFD Variants with High Carboligation Activity in the Presence of Water-Miscible Organic Solvents A basic problem concerning the application of hydrophobic aldehydes as substrates for the enzymatic formation of 2-hydroxy ketones is their low sol-


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ubility in aqueous buffer. A catalytic process in the presence of organic solvents is of interest to increase the solubility of the aldehyde substrates and to facilitate the recovery of products. However, enzymes that are optimized for biological systems often show low stability and catalytic activity in nonnatural environments (15). In the case of BFD, the wild-type enzyme is sufficiently stable in the presence of organic solvents; the best results with regard to increased solubility of hydrophobic substrates together with the least loss of carboligase activity of BFD were obtained by the addition of DMSO (5,7). In this way, (R)-benzoin (ee > 99%) was obtained in 70% yield (7). The reaction that shall be synthetically used is the side reaction of the enzyme, which is characterized by a significantly lower catalytic activity than the main reaction (decarboxylation). The application of enzymes in technical processes requires catalytic activities as high as possible to allow optimal yields, so BFD variants with enhanced carboligase activity in the presence of organic solvents were developed by directed evolution, which has rapidly emerged as the method of choice for the development and selection of mutated enzymes with improved properties (16). A mutant library was generated by error-prone PCR (17). Using a screening system in which the carboligase activity of the mutant enzymes was monitored by a rapid colorimetric assay (18), BFD with increased carboligase activity in aqueous buffer and mutein with enhanced carboligase activity in water-miscible organic solvents were generated. In the highly active mutein residue, Leu476, which is not located in the active center of the enzyme, was mutated. Leu476 turned out to be a hot-spot region for carboligase activity and has been investigated further by saturation mutagenesis, which was shown to enhance the effectiveness of directed evolution (19). This approach resulted in eight muteins with different residues at this position, with up to five-fold increased carboligase activity as compared to the BFD wild-type enzyme (Table 3). Surprisingly, all L476 mutein catalyze the formation of 2-HPP with significantly higher enantioselectivity than the wildtype enzyme, although enantioselectivity was not a selection parameter. Leucine 476 potentially plays the role of a gatekeeper of the active site of BFD, possibly by controlling the release of the product. An interaction of L476 with residues that support the V-conformation of the cofactor TDP, which is energetically unfavorable but common among the TDP-dependent enzymes, could be possible (20). The increased carboligation activity could be the result of a greater coenzyme flexibility. D. BFD Variants as Solution of the ‘‘ortho-Problem’’ Screening the same library with 2-methylbenzaldehyde and acetaldehyde as substrates yielded two BFD variants, BFD L476Q and BFD M365L-L461S, accepting ortho-substituted benzaldehyde derivatives as donor substrates.

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Lingen et al. Table 3 Initial Rate Velocities and Enantioselectivity of Wild-Type BFD and Muteins Concerning the Formation of (S)-2-HPPa

Carboligation activity (U/mg) Enzyme Wild-type BFD L476Q-S525G L476Q L476[Pub] L476S L476C L476M L476T L476A L476G L476H L476K

Buffer 7.1 17.4 28.1 4.6 20.6 19.2 27.1 23.3 17.3 16.8 14.4 20.7

F F F F F F F F F F F F

0.7 1.3 2.5 0.2 0.6 1.0 1.4 1.2 1.5 1.7 0.8 0.2

20% DMSO

eeb (%) [(S)-2-HPP]

F F F F F F F F F F F F

83 94 95 94 92 95 93 94 96 95 93 96

6.4 20.8 32.4 10.7 22.4 27.6 27.1 25.5 26.3 27.3 20.4 32.0

0.7 2.0 1.7 0.5 1.3 2.1 1.2 1.2 2.5 2.8 5.2 0.4

a 50 mM Kpi, pH 7.0; 2.5 mM MgSO4; 0.5 mM TDP; 40 mM benzaldehyde; 500 mM acetaldehyde; 30jC. b ee = enantiomeric excess. Source: Ref. 17.

Carboligation of these aldehydes could result in different products, including ortho-substituted benzoin or 2-HPP derivatives, depending on which aldehyde was accepted as donor and/or acceptor substrate. Both muteins, BFD L476Q and BFD M365L-L461S, catalyze the formation of enantiopure (S)-2hydroxy-1-(2-methylphenyl)propan-1-one [(S)-12c] with excellent conversion rates. Different ortho-substituted benzaldehyde derivatives, such as 2-chloro-, 2-methoxy-, and 2-bromobenzaldehyde, were accepted as donor substrates by both enzymes, and conversion with acetaldehyde resulted in the corresponding (S)-HPP derivatives 11a–c (Scheme 4) (21). III. DIFFERENT REACTIONS CATALYZED BY BENZALDEHYDE LYASE Benzaldehyde lyase (BAL, E.C. 4.1.2.38) from Pseudomonas fluorescens was first described by Gonza´les and Vicunã (22). They showed that this strain can grow on benzoin as a sole carbon and energy source, due to the ability of BAL


123

Scheme 4 BFD L476Q-mediated carboligation on a preparative scale of orthosubstituted benzaldehyde derivatives (7a–c) and acetaldehyde, yielding (S)-11a–c. (From Ref. 21.)

to catalyze the cleavage of the acyloin linkage of benzoin. When racemic benzoin was reacted with BAL (23) in potassium phosphate buffer, only a very small amount of benzaldehyde was formed. The addition of 20% DMSO as a cosolvent or alternatively 15% polyethylene glycol (PEG 400) resulted in enhanced benzaldehyde formation (24). Only (R)-benzoin is converted into benzaldehyde through BAL catalysis, although complete conversion of (R)benzoin was not possible under the conditions tested. Apparently, an equilibrium between cleavage and formation of (R)-benzoin exists during this process. (S)-benzoin gave no reaction at all. According to mechanistic considerations and assuming that cleavage and formation of (R)-benzoin are in equilibrium (24), BAL should also catalyze carboligation. Consequently, BAL-catalyzed acyloin condensation of benzaldehyde in aqueous buffer–DMSO mixtures resulted in almost quantitative formation of enantiomerically pure (R)-benzoin. The reaction was carried out on a preparative scale with different aromatic and heteroaromatic aldehydes (Table 4) (25). In contrast to wild-type BFD, BAL accepts aromatic aldehydes substituted at the ortho-position as well. Only a few aromatic aldehydes, such as pyridine 3- and 4-carbaldehyde, as well as the sterically exceedingly demanding aldehydes, gave either a very low yield or no benzoin condensation at all (25). A. Kinetic Racemic Resolution via C–C Bond Cleavage For nonenzymatic benzoin condensations it is well established that benzoins can be used instead of aldehydes as substrates. When (R)-benzoin was reacted with BAL in the presence of acetaldehyde, quantitative formation of enantiopure (R)-2-HPP occurred. The same reaction starting from (S)-benzoin failed. Repeating this reaction with racemic benzoin afforded enantiopure (R)-2-HPP and (S)-benzoin after separation of the products by column chromatography (Scheme 5) (24,25). Since the benzoin condensation cata-

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Lingen et al. Table 4 BAL-Catalyzed Benzoin Condensation on a Preparative Scale

Ar Ph 2-FC6H4 2-ClC6H4 2-MeOC6H4 3-FC6H4 3-ClC6H4 3-MeOC6H4 4-FC6H4 4-ClC6H4 4-MeOC6H4

Yield (%)

eea (%)

Configuration

96 68 80 87 80 94 93 89 95 95

>99 96 97 >99 97 >99 >99 >99 >99 >99

(R) (R) (R) (R) (R) (R) (R) (R) (R) (R)

a ee = enantiomeric excess. Source: Ref. 25.

lyzed by BAL gives access to the (R)-enantiomer of benzoins, either enantiomer of the respective benzoin can be obtained via the same enzyme using two different reactions. As expected from these results, the BAL-catalyzed reaction of benzaldehyde and acetaldehyde also gave (R)-2-HPP in 94% yield (Table 5). Different aromatic and heteroaromatic aldehydes are accepted as substrate for this carboligation reaction (24,25). Because there is still a lack of structural information about BAL, a structure-based discussion of the observed stereocontrol is not yet possible. Since BAL and BFD are enantiocomplementary with regard to 2-HPP formation (see Tables 1 and 5), most 2-HPP derivatives

Scheme 5 BAL-catalyzed kinetic racemic resolution via C–C bond cleavage. (From Ref. 24.)


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Table 5 BAL-Catalyzed Carboligation on a Preparative Scale Toward (R)-2-HPP derivatives

Ar Ph 2-FC6H4 2-MeOC6H4 3-ClC6H4 3-MeOC6H4 4-ClC6H4 4-MeOC6H4 3,5-di-FC6H3

Yield (%)

eea (%)

Configuration

94 64 63 94 80 88 64 67

>99 97 >99 >99 >99 >99 >99 >99

(R) (R) (R) (R) (R) (R) (R) (R)

a

ee = enantiomeric excess. Source: Refs. 24 and 25.

substituted in the ortho-, meta-, or para-position can be synthesized in either enantiomeric form using BFD and BAL wild-type enzymes and the newly developed BFD muteins (see Chapter 3). B. Asymmetric Cross-Benzoin Condensation with BAL As shown in Section II.B, the enzyme-catalyzed benzoin condensation starting from two different aromatic aldehydes can result in the selective formation of cross-benzoin products. The substrate spectrum of BAL in comparison to wild-type BFD with regard to the benzoin condensation is significantly broader, especially for sterically demanding substrates and benzaldehyde derivatives substituted in the ortho-position. Consequently, for BAL-catalyzed asymmetric cross benzoin condensation, a somewhat different set of selective donor and selective acceptor substrates is observed. A broad variety of unsymmetrical (R)-benzoins is accessible through BAL-catalyzed transformation, among which 12, 13, and 14 were obtained selectively, demonstrating that 3,4-methylendioxy-benzaldehyde, 3-hydroxy-4-methoxybenzaldehyde, and 3,4,5-trimethoxybenzaldehyde were selective donor substrates for BAL (11). Besides 2-chlorobenzaldehyde, fluorinated benzaldehyde derivatives proved to be selective acceptor substrates for BAL-catalyzed asymmetric cross-benzoin condensation (Scheme 6, compounds 15–17) (11). Direct access to the corresponding (S)-configured mixed benzoins is also given by use of BAL in a racemic resolution via C–C bond cleavage, a reaction

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Scheme 6 Mixed (R)-configured benzoins accessible through BAL-catalyzed asymmetric cross-benzoin condensation. (From Ref. 11.)

known from the synthesis of symmetrical (S)-benzoins (see Section III.A). In our investigation, racemic 2V-chloro-3,4,5-trimethoxy benzoin (rac-14) was reacted with an excess of acetaldehyde, which is a very potent acceptor molecule in the presence of BAL, to yield the mixed (S)-benzoin [(S)-15] in addition to (R)-1-(2-chlorophenyl)-2-hydroxy-propanone and (R)-2-hydroxy 1-(3,4,5-trimethoxyphenyl)propanone (11). Thus, we have shown that mixed benzoins can be synthesized enantioselectively through enzymatic cross benzoin condensation by TDP-dependent enzymes, taking advantage of the aldehydes’ donor–acceptor attitude. This one step synthesis starting from cheap and commercially available aldehydes represents an outstanding improvement in comparison to the costly and tedious synthesis based on the conversion of chiral cyanohydrins with phenyl-Grignard derivatives.

IV. SUMMARY BFD and BAL have been established as tools for enantioselective C–C bond formation, leading to enantiopure unsymmetrical and symmetrical 2-hydroxy ketones, which are important structural subunits in many biologically active natural products and are important building blocks for stereoselective synthesis (26). The detailed investigation of the TDP dependent enzymes BFD


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and BAL resulted in new concepts in chemoenzymatic synthesis. Wild-type BFD was shown to catalyze the formation of enantiopure meta- or parasubstituted (R)-benzoins and (S)-2-HPP derivatives (4–7). The enantioselectivity of the carboligation to HPP and derivatives is influenced by temperature, benzaldehyde concentration, and the electronic and steric properties of aromatic substituents of the donor aldehyde. Using rational protein design as well as directed evolution methods, together with substrate engineering, mixed benzoins could be synthesized enantioelectively through enzymatic cross-benzoin condensation (11), and BFD variants with enhanced carboligase activity in the presence of organic solvents (17). The potential of BAL for catalyzing C–C bond formation was investigated, leading to the synthesis of symmetrical (R)-benzoins and (R)-2-HPP derivatives with a broad substrate range (25). With BAL as catalyst, a kinetic resolution of racemates by C–C bond cleavage and concomitant C–C bond formation was established, giving access to both enantiomers of benzoins and (R)-2-HPP analogs (24). In this way, BAL, BFD, and BFD mutants complement one another, since BAL and BFD are enantiocomplementary with regard to 2-HPP formation. meta-, para- and ortho-substituted 2 HPP derivatives can be synthesized in either enantiomeric form using BAL, BFD, and BFD mutants.

REFERENCES 1. 1a. 1b. 1c. 1d. 2a.

2b.

3.

4.

For some recent reviews of a-keto acid decarboxylases and other TDP-dependent enzymes with regard to asymmetric synthesis, see: GA Sprenger, M Pohl. Synthetic potential of thiamine diphosphate-dependent enzymes. J Mol Catal B 6:145–159, 1999. U Scho¨rken, GA Sprenger. Thiamin-dependent enzymes as catalysts in chemoenzymatic syntheses. BBA 1385:229–243, 1998. H Iding, P Siegert, K Mesch, M Pohl. Application of a-keto acid decarboxylases in biotransformations. BBA 1385:307–322, 1998. OP Ward, A Singh. Enzymatic asymmetric synthesis by decarboxylases. Curr Opin Biotechnol 11:520–526, 2000. R Wilcocks, OP Ward, S Collins, NJ Dewdney, Y Hong, E Prosen. Acyloin formation by benzoylformate decarboxylase from Pseudomonas putida. Appl Environ Microbiol 58:1699–1704, 1992. R Wilcocks, OP Ward. Factors affecting 2-hydroxypropiophenone formation by benzoylformate decarboxylase from Pseudomonas putida. Biotechnol Bioeng 39:1058–1063, 1992. H Iding, Struktur- und Funktionsuntersuchungen an der Benzoylformiatdecarboxylase aus Pseudomonas putida. PhD dissertation, Heinrich Heine University, Du¨sseldorf, Germany, 1998. H Iding, T Du¨nnwald, L Greiner, A Liese, M Mu¨ller, P Siegert, J Gro¨tzinger, AS Demir, M Pohl. Benzoylformate decarboxylase from Pseudomonas putida as

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6. 7.

8.

9. 10. 11.

12.

13a. 13b.

13c. 13d. 13e. 14a. 14b. 15. 16. 17.

Lingen et al. stable catalyst for the synthesis of chiral 2-hydroxy ketones. Chem Eur J 6:1483– 1495, 2000. T Du¨nnwald, AS Demir, P Siegert, M Pohl, M Mu¨ller. Enantioselective synthesis of (S)-2-hydroxypropanone derivatives by benzoylformate decarboxylase catalyzed C–C bond formation. Eur J Org Chem 11:2161–2170, 2000. T Du¨nnwald, M Mu¨ller. Stereoselective formation of bis(a-hydroxy ketones) via enzymatic carboligation. J Org Chem 65:8608–8612, 2000. AS Demir, T Du¨nnwald, H Iding, M Pohl, M Mu¨ller. Asymmetric benzoin reaction catalyzed by benzoylformate decarboxylase. Tetrahedron: Asymmetry 10:4769–4774, 1999. MS Hasson, A Muscate, MJ McLeish, LS Polovnikova, JA Gerlt, GL Kenyon, GA Petsko, D Ringe. The crystal structure of benzoylformate decarboxylase at 1.6-A˚ resolution: diversity of catalytic residues in thiamin diphosphate– dependent enzymes. Biochemistry 37:9918–9930, 1998. LP Hammett. The effect of structure upon the reactions of organic compounds. Benzene derivatives. J Am Chem Soc 59:96–103, 1937. D Kihumbu, T Du¨nnwald, M Mu¨ller, J Bargon, A Liese. To be published. P Du¨nkelmann, D Kolter-Jung, A Nitsche, AS Demir, P Siegert, B Lingen, M Baumann, M Pohl, M Mu¨ller. Development of a donor–acceptor concept for catalytic asymmetric cross-coupling reactions of aldehydes: the first asymmetric cross-benzoin condensation. J Am Chem Soc 124:12084–12085, 2002. LS Polovnikova, MJ McLeish, EA Sergienko, JT Burgner, NL Anderson, AK Bera, F Jordan, GL Kenyon, MS Hasson. Biochemistry 42:1820–1830, 2003. We thank Dr. McLeish for kindly providing us with the BFD H281A gene. E Fischer. Ueber das Furfural. Annalen Chemie 211:214–232, 1882. H Staudinger. U¨ber die Autoxydation organischer Verbindungen. II. Beziehungen zwischen Autoxydation und Benzoin-Bildung. Berichte der deutschen chemischen Gesellschaft 43:3535–3538. 1913. JS Buck, WS Ide. Mixed benzoins. I. J Am Chem Soc 52:220–224, 1930. G Semerano. Proprieta´ ossido-riduttive delle aldeidi e condensazione benzoinica. Gazz Chem Ital 71:447–461, 1941. KW Merz, D. Plauth. Benzoinkondensationen mit 4-acetylamino-benzaldehyd. Chemische Berichte. 90:1744–1757, 1957. JET Corrie. Preparation and properties of unsymmetrical benzoins and related compounds. Tetrahedron 54:5407–5415, 1998. MD Rozwadowska. Cyanohydrins as substrates in benzoin condensation; regiocontrolled mixed benzoin condensation. Tetrahedron 41:3135-3140, 1985. FH Arnold. Protein engineering for unusual environments. Curr Opin Biotechnol 4:450–455, 1993. UT Bornscheuer, M Pohl. Improved biocatalysts by directed evolution and rational protein design. Curr Opin Chem Biol 5:137–143, 2001. B Lingen, J Gro¨tzinger, D Kolter, MR Kula, M Pohl. Improving the carboligase activity of benzoylformate decarboxylase from Pseudomonas putida by a combination of directed evolution and site-directed mutagenesis. Protein Eng 15:585–593, 2002.


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18. M Breuer, M Pohl, B Hauer, B Lingen. Anal Biomed Chem 374:1069–1073, 2002. 19a. K Miyazaki, FH Arnold. Exploring nonnatural evolutionary pathways by saturation mutagenesis: rapid improvement of protein function. J Mol Evol 49:716–720, 1999. 19b. K Liebeton, A Zonta, K Schimossek, M Nardini, D Lang, BW Dijkstra, MT Reetz, KE Jaeger. Directed evolution of an enantioselective lipase. Chemistry Biol 7:709–718, 2000. 20. F Guo, D Zhang, A Kahyaoglu, RS Farid, F Jordan. Is a hydrophobic amino acid required to maintain the reactive V-conformation of thiamin at the active center of thiamin diphosphate-requiring enzymes? Experimental and computational studies of isoleucine 415 of yeast pyruvate decarboxylase. Biochemistry 37:13379–13391, 1998. 21. B Lingen, D Kolter, P Du¨nkelmann, R Feldmann, J Gro¨tzinger, M Pohl, M Mu¨ller. Alteration of the substrate specificity of benzoylformate decarboxylase from Ps. putida by directed evolution. CHEMBIOCHEM 4:721–726, 2003. 22a. B Gonza´lez, R Vicunã. Benzaldehyde lyase, a novel thiamine PPi-requiring enzyme, from Pseudomonas fluorescens biovar I. J Bacteriol 171:2401–2405, 1989. 22b. P Hinrichsen, I Go´mez, R Vicunã. Cloning and sequencing of the gene encoding benzaldehyde lyase from Pseudomonas fluorescens biovar I. Gene 144:137– 138, 1994. 23. The enzyme used in our studies was expressed and purified from a recombinant Escherichia coli strain. For easier purification, a hexahistidine tag was cloned to the C-terminus of the enzyme; E Janzen, M Pohl, unpublished. 24. AS Demir, M Pohl, E Janzen, M Mu¨ller. Enantioselective synthesis of hydroxy ketones through cleavage and formation of acyloin linkage. Enzymatic kinetic resolution via C–C bond cleavage. J Chem Soc, Perkin Trans 1 7:633–635, 2001. 25. AS Demir, O¨sesenoglu, E Eren, B Hosrik, M Pohl, E Janzen, D Kolter, R Feldmann, P Du¨nkelmann, M Mu¨ller. Enantioselective synthesis of a-hydroxy ketones via benzaldehyde lyase-catalyzed C–C bond formation reaction. Adv Synth Catal 344:96–103, 2002. 26. Cf. Ref. 25 and references cited therein.

9 Benzoylformate Decarboxylase: Lessons in Enzymology Michael J. McLeish and George L. Kenyon University of Michigan, Ann Arbor, Michigan, U.S.A. Elena S. Polovnikova*, Asim K. Bera, Natalie L. Anderson, and Miriam S. Hasson Purdue University–West Lafayette, West Lafayette, Indiana, U.S.A.

I. INTRODUCTION BFD is as an example of recent evolution. It is a member of the mandelate catabolic pathway, present only in Pseudomonas putida and a few closely related species. Collaborative studies on this pathway, begun by George Kenyon, Gregory Petsko, John Gerlt, Dagmar Ringe, and John Kozarich, have provided a rich understanding of the individual component enzymes (1). The pathway is composed of at least five unrelated enzymes, each evidently recruited from a different enzyme family. Two well-studied members of this pathway are mandelate racemase and mandelate dehydrogenase. Our investigation of BFD, a third member of the pathway, has provided answers and posed new questions regarding enzymes that use thiamine diphosphate (TDP) as a cofactor. Structural and biochemical comparison of BFD with other thiamine diphosphate (TDP)–dependent enzymes has increased our understanding of *Current affiliation: Kilpatrick Stockton LLD, Atlanta, Georgia, U.S.A.

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the forces shaping enzyme evolution (2). The overall architecture of BFD closely resembles that of related enzymes, and residues that bind the cofactor or metal ion are extremely well conserved. Surprisingly, none of the active-site residues (except those directly bound to cofactor) are conserved between any of the TDP-dependent enzymes. Even the two decarboxylases, BFD and pyruvate decarboxylase (PDC), have quite distinct active-site architecture. This represents a challenge to current expectations and understanding of enzyme evolution. Biochemical comparison of BFD- and PDC-catalyzed reactions also indicates that there are differences in the mechanism of catalysis of the later reaction steps. While catalytic residues differ, there remains a positional conservation of catalytic groups relative to the cofactor and the protein scaffold. These comparisons suggest that cofactor chemistry, the nature of the reaction intermediates, and architectural considerations relating to the protein fold have been dominant forces in the evolution of TDPdependent enzymes. The rationalization of the differences between the active sites requires elucidation of the roles of alternative catalytic residues in these enzymes. This will allow a more meaningful comparison of active-site structures and an understanding of the relationship between catalytic chemistry and structural evolution. We describe here our study using a combination of crystallographic structure determination, site-directed mutagenesis, kinetic analysis, and protein chemistry. II. STRUCTURAL GIFTS OF BFD Insights into the steps of an enzymatic reaction can be disclosed in many ways. For example, in crystallography, time-resolved studies are very enlightening. Common approaches focus on trapping the intermediates of the reaction using chemical or temporal methods. Yet in the study of BFD, favorable properties of the protein itself allow the growth of many crystal forms with different characteristics under similar conditions (Table 1). These crystals, grown in the same drop, are in different space groups and, most importantly, have different binding properties. The first published structure of BFD was determined from crystals in form I, showing a water molecule bound in the active site with hydrogen bonds to C2 and N1V of TDP and to the side chains of Ser26 and His70 (Fig. 1A). Form I is the most accessible known crystal form for solving the structure, because the crystals grow quickly and diffract very well, allowing the original structure to be solved to 1.6 A˚ in 1994 (2). The other crystal forms we have studied, containing 4 or 16 monomers in the asymmetric unit, have exposed other characteristics of BFD, many of which were neither predicted nor foreseen. In all the solved crystal structures of BFD the main contacts between the tetramers are mediated by a 13-residue polyproline type II (ppII) helix

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Table 1 Comparison of BFD Crystal Forms Crystal A b c a form (A˚) (A˚) (A˚) (j)

h (j)

g (j)

Space group

Highest resolution (A˚)

I

82

97 138

90

90

90 I222

1.0

II

94 113 205

90

90

90 P21212

2.8

90 90 90 64 90

90 90 97 72 90

90 90 90 73 90

— — 3.3 1.3 2.0

III IV V VI VII

107 98 136 70 70

159 101 210 92 160

280 110 164 94 176

P21212 I222 P21 P1 P212121

Well solution A (22% PEG 400, 0.15 M CaCl2, 0.5% MPD, Tris-Cl pH 8.5) B (20–28% PEG-MME 2000, 0.15–0.3 M (NH4)2SO4, 0.1 M Na-citrate pH 5–5.6) B B B A A

Monomers/ ASU 1

4

— — 16 4 4

Figure 1 Structures of the BFD active site in crystal form I (A) and in crystal form V with (R)-mandelate bound (B).

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(Fig. 2). This element of secondary structure is quite common in globular proteins, but it is unusual for the helix to be composed of more than five residues (3). The connection between the h and g domains of PDC also forms a ppII helix. Conserved ppII helices 10–13 residues in length also occur in molecules that interact with SH3 domains (4,5) and peptides bound to major histocompatibility complex molecules for presentation to T cells (6–8). The ppII helices mediate important protein/protein interaction in both cases. We have proposed that, in the same way, the appearance of crystal contacts through the ppII helix in BFD suggests that the helix may also be involved in protein/protein interactions in the bacterial cell (2). This proposal has been strengthened by the involvement of the ppII helix in contacts in all of the four crystal forms of BFD that have been solved [(9,10); work in progress]. In vivo, BFD is evidently part of a multienzyme complex that includes other members of the mandelate pathway (11). Perhaps the long, regular ppII helix, exposed on the surface of BFD, is a convenient handle by which BFD is held in the complex.

Figure 2 The main crystal contact between tetramers in crystal form I. The polyproline type II helix is involved in crystal contact in all solved crystal structures of BFD.


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III. ENZYMATIC ROLES OF BFD RESIDUES A generally accepted model of the reaction catalyzed by BFD is shown in Figure 3. The reaction begins with the deprotonation of C2 by the N4V-imino group of TDP and binding of the substrate, benzoylformate. The first intermediate, 2-a-mandelyl-TDP (M-TDP), is formed by attack of the C2 ylide on the carbonyl group of the substrate. Decarboxylation of M-TDP forms the second intermediate, the 2-a-carbanion, or enamine. The enamine

Figure 3 The reaction catalyzed by BFD, illustrating possible roles of active-site residues.

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is then protonated, forming 2-a-hydroxylbenzyl-TDP (HB-TDP), which then breaks down to release benzaldehyde. A. Structural Studies We have examined the roles of the active-site residues through molecular biological, kinetic, and structural studies. Site-directed mutagenesis and kinetic studies suggested that four residues in the active site that are not in direct contact with the cofactor TDP, Ser26, Glu28, His70, and His281, all have a strong effect on the reaction. To understand the roles of these residues, a structure of the enzyme binding a substrate or inhibitor was essential. The first structure we solved contained a water molecule near C2 of TDP, the site where the substrate would be expected to bind. More recently we have obtained crystal form V (Table 1), grown at pH 5.5, in which BFD was liganded to (R)-mandelate, an inhibitor in which the keto group of the substrate has been reduced (Fig. 1B). An interesting aspect of the binding of the (R)-mandelate is the fact that not only are all of its hydrophilic atoms involved in hydrogen bonds, as expected, but that each of these atoms is also surrounded by hydrophobic residues located 3–3.5 A˚ away (Fig. 4). The side chains of Leu110, Leu461, and Phe464 have close contacts with the hydroxyl and carboxylate groups. These hydrophobic residues may assist by exclusion of water molecules from the reaction center, thereby reducing side reactions. Intriguingly, considering the hydrophobic portion of the substrate, it might have been expected that stacking interactions between the phenyl ring of the substrate and the aromatic side chains in the active site would play a role in substrate binding. This is especially true because only benzoylformate and its para-substituted analogs can serve as BFD substrates (12). However, only two aromatic residues, Phe397 and Phe464, are within 4 A˚ of R-mandelate, and their rings do not have the expected stacking interactions. The structure suggests key roles for the active-site residues under study. One nitrogen group in the His70 ring is within hydrogen-binding distance from the hydroxyl group of (R)-mandelate suggesting that this residue is involved in the protonation and deprotonation of the carbonyl group of benzoylformate. The second nitrogen in the His70 ring forms a hydrogen bond with the carboxyl group of Glu28, suggesting a possible proton relay. Ser26 forms hydrogen bonds with the carboxylate group of the inhibitor, implicating this residue in substrate binding and possibly suggesting the role for this residue in promoting decarboxylation. His281 is somewhat removed from the inhibitor, making it difficult to predict its role in the reaction. However, (R)-mandelate is not in the identical configuration to the substrate, because the carbon attacked by C2 of TDP is sp3 rather than sp2


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Figure 4 Binding of R-mandelate to the active site in crystal form V. Dashed lines signify contacts V3.0 A˚ (dark) and other contacts V4.0 A˚ (light).

hybridized. It is likely that the conformation of the active site changes as the reaction proceeds, in terms of both side chains and main-chain movements, some of which were observed in this structure (Phe464, for example; see Fig. 4). In addition, while the first and third domains both contribute to the active site of BFD, His 281 is a member of the second domain. This may allow H281 more freedom to move relative to the other residues in the active site, permitting a role that may not be obvious from the X-ray structures alone.

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B. Kinetic Studies The structural picture of (R)-mandelate-bound BFD dovetails with kinetic studies on site-directed mutants of the protein. Steady-state kinetic studies indicated that the catalytic parameters are strongly affected in Ser26Ala, His70Ala, and His281Ala (10). The Km value for benzoylformate and the Ki value for R-mandelate were affected to the highest degree in the Ser26Ala mutant, increasing 25-fold and 100-fold, respectively. This observation agrees with the structural analysis, suggesting that the serine hydroxyl is important for substrate binding. The kcat value for H70A is reduced by more than three orders of magnitude, strongly implicating this residue in catalysis, correlating with its predicted role of protonation and deprotonation of the intermediates. His281 showed significant, but diminished, effects on both Km and kcat, consistent with a lesser role for this residue. With the group of Frank Jordan at Rutgers, we have further analyzed the roles of the residues through stopped-flow spectroscopy (10,13). Using the substrate ( p)-nitrobenzoylformate, the rates of the production and decay of the M-TDP and enamine intermediates appear to be reflected in the changes in absorbance at 620 nm and 420 nm, respectively. The biphasic breakdown of the second peak (at 420 nm) suggests that a slow, reversible protonation of the enamine that allows the observation of the nonreversible breakdown of the next intermediate, HB-TDP, as well. Using this interpretation of the wild-type curves, stopped-flow experiments using His70Ala, Glu28Ala, His281Ala, and Ser26Ala mutants suggest roles very similar to those proposed by examination of the structure with (R)mandelate bound. His70 and Glu28 are involved in the protonation of the substrate to form M-TDP and deprotonation of HB-TDP to form the product. The enamine, the most stable intermediate, requires some destabilization to continue on in the reaction; apparently, His281 is involved in this process. Surprisingly, the data collected on the Ser26Ala mutant suggest that Ser26 is involved not only in the removal of the carboxylate group but in later steps as well. Our structural results suggest that Glu28 may play a role in mediating the pKa of His70 as well as the positioning of the imidazole ring (manuscript in preparation). The results of kinetic studies (9) have supported this notion. In a structure of the Glu28Ala mutant, an oxygen atom of a water molecule replaces the one of the oxygen atoms in the carboxylate group of Glu28 and forms a hydrogen bond with the nitrogen of the His70 side chain. At low pH, the Glu28Ala mutant is catalytically active, as observed by both steady-state and stopped-flow studies, but at higher pH the enzyme becomes significantly less active than WT. His70 is thought to play a role as a general acid, providing a proton to the carbonyl group of benzoylformate, thereby


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facilitating the production of M-TDP (Fig. 3). It is likely that Glu28 raises the pKa of His70, allowing it to be active over a broad pH range. In the absence of Glu28, the enzyme is active at low pH, where His70 would be expected to be protonated. However, as the pH rises above 7.0, the histidine is deprotonated and the enzyme rapidly becomes less active. In summary, given the structural and kinetic data described up to this point, we can suggest the roles of four residues in the active site of BFD. His70, with the help of Glu28, assists in the transfer of protons in the first and last steps of the reaction. His281 is involved in the protonation of the enamine. Ser26 binds the carboxylate group of the substrate, and also plays a later role in the reaction. IV. AN UNEXPECTED GUEST IN THE ACTIVE SITE In crystals of wild-type BFD in crystal form I, only a water molecule has been found in the active site, even when substrate or inhibitors have been soaked into the crystal. However, in crystals of form VI, crystallized under the same conditions and often present in the same drop as crystal form I, bicarbonate (or carbonate) appears, forming hydrogen bonds with Ser26, His70, His281, and TDP (manuscript in preparation). There has not been much thought given to the fate of the atoms of the carboxylate group that is removed in TDP-dependent decarboxylation. Normally, the carbon dioxide released in the decarboxylation step is thought to diffuse away, with no further effect on the reaction. The discovery of bicarbonate in the active site of BFD and carbon dioxide in pyruvate:ferrodoxin oxidoreductase (14) suggests that this product may play some later role in the reaction. As one possibility of many, we suggest that the ability of the enzyme to bind bicarbonate may increase its ability to release its second product, benzaldehyde. V. THE POWER OF Ser26 Our results suggest that Ser26 has a role in the later steps of the reaction, after decarboxylation has occurred (10). The indication is somewhat difficult to understand. The evidence suggests that Ser26 helps bind the substrate and facilitates decarboxylation. Stopped-flow studies have also suggested that Ser26 plays a role in deprotonation of HB-TDP and release of benzaldehyde. It is not obvious to us how Ser26, located close to the carboxylate group but not to the rest of the substrate or other intermediates, can have an effect after carbon dioxide has been released. Further, the unexpected results of an experiment we have done with Theodore Widlanski at Indiana University have also suggested that Ser26 is somewhat unusual. When BFD was

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incubated with benzoylphosphonate, both analysis of a structure solved in crystal in form II and mass spectrometric analysis indicated that Ser26 was phosphorylated (9). Prior to this, to the best of our knowledge, phosphorylation of a serine by a phosphonate has not been observed in enzymology. Several of our recently solved structures show that the Ser26 hydroxyl group is able to swing around, away from the active site. It is possible that the CO2 eliminated during the decarboxylation step may be removed from the active site through the movement of the serine, either as CO2, as in yeast PDC (15), or as bicarbonate. In BFD product release is rate limiting, and it is conceivable that the removal of CO2 from the active site or, possibly, the return of bicarbonate to the active site assists in the release of benzaldehyde. If that were the case, the role of the Ser26 in controlling the movement and the hydration of carbon dioxide would explain its importance throughout the entire reaction. This is of course a hypothesis; we are currently undertaking experiments to help understand the unusual results.

VI. CONCLUSIONS BFD has indeed given us valuable lessons on the nature of the TDP-dependent enzyme family. We hope those aspects we do not yet understand and those we have not yet imagined will be revealed in our further studies.

ACKNOWLEDGMENTS This research was supported at Purdue by NSF grant 9733552-MCB award and by David and Lucille Packard Foundation Fellowship 99-1463 (to MSH) and by USPHS-NIH-GM-40570 (to GLK). Facilities shared by the Structural Biology group at Purdue have been developed and supported by grants from NIH, NSF, the Lucille P. Markey Foundation, the Keck Foundation, and the office of the university executive vice president for academic affairs at Purdue University.

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GA Petsko, GL Kenyon, JA Gerlt, D Ringe, JW Kozarich. On the origin of enzymatic species. Trends Biochem Sci 18:372–376, 1993. MS Hasson, A Muskate, MJ McLeish, LS Polovnikova, JA Gerlt, GL Kenyon, GA Petsko, D Ringe. The crystal structure of benzoylformate decarboxylase at 1.6-A˚ resolution: diversity of catalytic residues in thiamin diphosphate– dependent enzymes. Biochemistry 37:9918–9930, 1998. AA Adzhubei, MJ Sternberg. Conservation of polyproline II helices in


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homologous proteins: implications for structure prediction by model building. Protein Sci 3:2395–2410, 1994. GB Cohen, R Ren, D Baltimore. Modular binding domains in signal transduction proteins. Cell 80:237–248, 1995. T Pawson. Protein modules and signalling networks. Nature 373:573–580, 1995. TS Jardetzky, JH Brown, JC Gorga, LJ Stern, RG Urban, JL Strominger, DC Wiley. Crystallographic analysis of endogenous peptides associated with HLADR1 suggests a common, polyproline II-like conformation for bound peptides. Proc Natl Acad Sci USA 93:734–738, 1996. LJ Stern, JH Brown, TS Jardetzky, JC Gorga, RG Urban, JL Strominger, DC Wiley. Crystal structure of the human class II MHC protein HLA-DR1 complexed with an influenza virus peptide. Nature 368:215–221, 1994. DR Madden, JC Gorga, JL Strominger, DC Wiley. The three-dimensional structure of HLA-B27 at 2.1-A˚ resolution suggests a general mechanism for tight peptide binding to MHC. Cell 70:1035–1048, 1992. LS Polovnikova. PhD dissertation, Purdue University, West Lafayette, IN. Supervised by M.S. Hasson. 2000. LS Polovnikova, MJ McLeish, EA Sergienko, JT Burgner, NL Anderson, AK Jordan, F Jordan, GL Kenyon, MS Hasson. Structural and kinetic analysis of catalysis by a thiamin diphosphate–dependent enzyme, benzoylformate decarboxylase. Biochemistry, in press, 2003. RA Halpin, GD Hegeman, GL Kenyon. Carbon-13 nuclear magnetic resonance studies of mandelate metabolism in whole bacterial cells and in isolated, in vivo cross-linked enzyme complexes. Biochemistry 20:1525–1533, 1981. LJ Reynolds, GA Garcia, JW Kozarich, GL Kenyon. Differential reactivity in the processing of [ p-(halomethyl)benzoyl] formates by benzoylformate decarboxylase, a thiamin pyrophosphate–dependent enzyme. Biochemistry 27:5530– 5538, 1988. EA Sergienko, J Wang, L Polovnikova, MS Hasson, M McLeish, GL Kenyon, F Jordan. Spectroscopic detection of transient thiamin diphosphate–bound intermediates on benzoylformate decarboxylase. Biochemistry 39:13862–13869, 2000. E Chabriere, X Vernede, B Guigliarelli, MH Charon, EC Hatchikian, JC Fontecilla-Camps. Crystal structure of the free radical intermediate of pyruvate:ferredoxin oxidoreductase. Science 294:2559–2563, 2001. M Liu, EA Sergienko, F Guo, J Wang, K Tittmann, G Hubner, W Furey, F Jordan. Catalytic acid–base groups in yeast pyruvate decarboxylase. 1. Sitedirected mutagenesis and steady-state kinetic studies on the enzyme with the D28A, H114F, H115F, and E477Q substitutions. Biochemistry 40:7355–7368, 2001.

10 New Concept on the Nature of the Induced Absorption Band of Holotransketolase Marina V. Kovina, Irina A. Sevostyanova, Olga N. Solovjeva, Ludmilla E. Meshalkina, and German A. Kochetov Moscow State University, Moscow, Russia

It has long been known that formation of a catalytically active holotransketolase from apoenzyme and thiamine diphosphate (TDP) is accompanied by the appearance, in both the absorption and CD spectra, of a new band that was lacking in the initial components. Binding and subsequent conversion of transketolase substrates bring about changes in this band’s intensity. The observation of these changes allows one to monitor the coenzyme-to-apoenzyme binding and the substrate’s conversion during the transketolase reaction and thus to kinetically characterize its individual steps. As regards the new absorption band, induced by TDP binding, its nature, until recently, remained unknown. The reason of its appearance was considered to be either the formation of charge transfer complex between TDP and tryptophan (phenylalanine) residue or stacking interaction between the residues of aromatic amino acids. They are thought to be brought together as a result of conformational changes of the apoenzyme during its interaction with the coenzyme. However, none of these hypotheses was substantiated experimentally. In our hypothesis, the induced absorption band is that of the imino form of TDP resulting from three contributing features of the coenzyme-binding site of transketolase. These factors are the relative hydrophobicity of this site, 143

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hydrogen bonding of the N1V atom of the TDP aminopyrimidine ring to Glu418 and base stacking interactions between the iminopyrimidine ring of TDP and Phe445. The TDP-dependent enzyme transketolase (EC 2.2.1.1) catalyzes the cleavage of a carbon–carbon bond and reversible transfer of a two-carbon unit (a,h-dihydroxyethyl group) from ketose, the donor substrate, to aldose, the acceptor substrate (1) (Scheme 1). Transketolase has a broad substrate specificity and can accept a variety of ketose sugars (xylulose 5-phosphate, fructose 6-phosphate, sedoheptulose 7-phosphate, erythrulose, etc.) and also h-hydroxypyruvic acid (HPA) as donor substrates. The use of HPA is of synthetic significance because it allows a practically irreversible product formation, with carbon dioxide leaving the assay. Ribose 5-phosphate, erythrose 4-phosphate, glyceraldehyde 3-phosphate, glycolaldehyde, etc. serve as the acceptor substrates. Transketolase from the yeast Saccharomyces cerevisiae is a well-investigated enzyme with a known three-dimensional structure and a general catalytic mechanism. A great deal of X-ray data have been collected on the 3D structures of the apo- and holoenzyme (2–5) as well as on various transketolase complexes with TDP analogs (6,7), a holotransketolase complex with the acceptor substrate (8), and of the key intermediate, a-carbanion of a, h-dihydroxyethyl-TDP (DHETDP) (9). The combination of site-directed mutagenesis, X-ray structural studies, the reaction kinetics, and CD spectroscopy data enabled the characterization of the amino acid residues involved in the interactions with TDP, in substrate channel formation and in catalysis (5,10–14). Transketolase is composed of two identical subunits, each with a molecular mass of 74 kDa, and has two active centers (15–17). Native holoenzyme contains Ca2+ (18), but other metal ions, such as Mg2+, Mn2+, and Co2+, can also serve as cofactors (19). TDP is bound at the interface between the subunits and interacts with residues from both subunits, and the dimer

Scheme 1 Transketolase reaction.

Induced Absorption Band of Holotransketolase

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may be considered as a catalytically competent unit (3). The comparison of the 3D structures of some thiamine diphosphate enzymes has shown that the so-called V-conformation of the TDP molecule provides a direct contact between the amino group of the pyrimidine ring and the C2–H bond of the thiazolium ring (20). TDP is almost completely buried inside the protein. The only atom of cofactor accessible from the bulk solution is the C2 atom of the thiazolium ring. The aminopyrimidine ring of TDP is located in a hydrophobic pocket formed particularly by the aromatic side chains, Phe442, Phe445, and Tyr448 (Fig. 1). The side chain of Phe445 is in a stacking interaction with the aminopyrimidine ring. Additionally, the aminopyrimidine ring forms several hydrogen bonds, the most significant of which is the one between

Figure 1 The hydrophobic ‘‘pocket’’ of the aminopyrimidine ring of TDP in holotransketolase (atomic coordinates are taken from the Protein Data Bank, ID = 1 Trk).

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the N1V-atom of the aminopyrimidine ring and the side chain of Glu418 (3, 5,10). Analogous interaction is observed with other TDP-dependent enzymes, which plays an important role in the mechanism of thiamine catalysis (21–23). The transketolase reaction can be divided into two parts. The first half of the reaction consists of the following steps: binding of donor substrate and formation of a covalent enzyme–substrate complex, cleavage of the donor substrate, and formation of a covalently bound key intermediate, DHETDP, and the first product, an aldose. In the second half of the reaction the intermediate interacts with the acceptor–aldose substrate. The two-carbon unit is transferred to the acceptor substrate, and the new ketose, with its carbon chain extended by two carbon atoms, is released. The interaction of apotransketolase with TDP and the formation of catalytically active holoenzyme is accompanied by the appearance of a new band in the absorption spectrum (range 290–350 nm, Fig. 2A, curve 1) and in the CD spectrum (range 300–380 nm, Fig. 2B, curve 2), which was lacking in the initial components (24,25). There is a clear-cut correlation between the quantity of TDP bound to the apoenzyme and the catalytic activity (17,26). This optical effect is widely used in experimental practice to investigate the process of TDP-to-apotransketolase binding (17,27). Figure 3 presents the equilibrium formation of holotransketolase as a function of TDP concentration in the presence of Ca2+ (curve 1) and Mg2+ (curve 2); the inset shows the initial part of the curves. Upon substitution of Ca2+ for Mg2+, the affinity of TDP to apotransketolase was markedly changed. The appearance of the new absorption band is characteristic of a catalytically active holoenzyme; its intensity would change after transketolase interaction with its substrates. The addition of the donor substrate (HPA) to holotransketolase leads to the formation of the key reaction intermediate, DHETDP. Formation of the intermediate is accompanied by the increase of the new band amplitude in the absorption spectrum (Fig. 2A, curve 2) and the inversion of the absorption band in the CD spectrum (Fig. 2B, curve 3). Subsequent addition of the acceptor substrate (glycolaldehyde) leads to the two-carbon unit transfer from intermediate to the acceptor substrate. This leads to formation of the second transketolase reaction product and the restoration of the initial holoenzyme and the holoenzyme CD spectrum (Fig. 2B, curve 4). So the interaction of the substrates with holotransketolase and their subsequent conversion is accompanied by significant changes in the intensity of the new absorption band, and these changes are widely used for characterization of the individual steps of the transketolase reaction (28–31). Transketolase is a typical transferase enzyme, requiring two substrates for catalysis, the two-carbon unit donor and acceptor, respectively. But


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Figure 2 (A) Difference absorption spectra of holotransketolase in the absence (1) and presence (2) of HPA with respect to apotransketolase, TDP, and HPA. (B) CD spectra of transketolase with TDP and substrates: 1—apotransketolase; 2—holotransketolase; 3—holotransketolase + HPA; 4—holotransketolase + HPA + glycolaldehyde. Glycyl-glycine buffer, 50 mM, pH 7.6; CaCl2, 2.5 mM; TDP, 40 lM; HPA, 2 mM; glycolaldehyde, 20 mM; transketolase, 3 lM (A) and 6 lM (B). Path length—1.0 cm.

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Figure 3 Equilibrium formation of holotransketolase as a function of TDP concentration. Glycyl-glycine buffer, 50 mM, pH 7.6; CaCl2 or MgCl2, 2.5 mM; transketolase, 6 lM.

surprisingly, the initial holoenzyme CD spectrum was restored even in the absence of acceptor substrate, although the restoration was not complete and occurred at a much slower rate, showing that the splitting of a two-carbon unit from the intermediate with the formation of free holoenzyme may occur in the absence of the acceptor substrate (29,30). This means that transketolase, being a typical transferase enzyme, is able to catalyze not only the common two-substrate reaction but also the cleavage of donor substrates, the first half-reaction, even in the absence of acceptor substrate. Conversion of the donor substrate HPA can be followed by using changes in the absorbance at 300 nm via stopped-flow kinetics (Fig. 4). The progress curve was followed under single-turnover conditions (equal concentration of HPA and the holotransketolase active sites). At least two phases could be observed: The increase of the absorption represents formation of the key intermediate DHETDP, decrease–release of the product, glycolaldehyde, and restoration of the holoenzyme. Each part of the progress curve can be fitted to a single exponential first-order reaction (data not shown). The rate


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Figure 4 Single turnover of HPA conversion by holotransketolase. Glycyl-glycine buffer, 25 mM, pH 7.6; CaCl2, 2.0 mM; TDP, 50 lM; transketolase, 6.7 lM (13.4 lM active site); HPA, 13.4 lM.

constants for the DHETDP formation and glycolaldehyde release differ about 30-fold. Thus the decomposition of DHETDP is the rate-limiting step under the experimental conditions. Although the new absorption band of holotransketolase was discovered a long time ago, its nature has not yet been identified. Initially it was attributed to the formation of a charge transfer complex between TDP and a tryptophan residue in the transketolase active center (32). However, according to X-ray data, the tryptophan residue closest to TDP is 19 A˚ away from it (data are taken from the Protein Data Bank, ID = 1 Trk). When the 3D structure of holotransketolase was solved, the tryptophan function was attributed to the Phe445 residue, which is in stacking interaction with the aminopyrimidine ring of TDP (5,10) (Fig. 1). To verify this suggestion, Phe445 was replaced by isoleucine. This replacement maintains the hydrophobic environment around the pyrimidine ring, but the aromatic ring disappears. The CD spectrum of mutant F445I has shown that this replacement does not lead to the disappearance of the induced absorption band, but is accompanied by a fivefold decrease in its intensity (33). Another explanation was then offered. As is known, binding of the TDP to the transketolase active site, followed by the conformational changes in the

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protein molecule, leads to the stabilization of two loops, which are flexible in apotransketolase and well ordered in holotransketolase (4). As a result, the Trp391 residue from the one loop and the Tyr370 residue from the other are brought closer together while their aromatic rings get parallel to each other. Based on this finding, it was suggested that the new, TDP-induced, absorption band is caused by stacking interaction between the aromatic rings of the two residues (33). To ascertain if this is indeed the case, Trp391 was replaced by phenylalanine. However, the CD spectrum of the mutant holotransketolase thus obtained proved very similar to that of wild-type holotransketolase (C. Wikner, personal communication, 1997). Thus none of the suggestions proposed to elucidate the nature of the induced absorption band whose appearance accompanies the TDP–apotransketolase interaction and the catalytically active holoenzyme formation appears plausible. In this chapter, a new hypothesis for the appearance of the induced absorption band of holotransketolase is proposed, according to which this phenomenon is caused by changed optical properties of TDP following its incorporation into the hydrophobic pocket of the enzyme’s active center. By now, the idea about the participation of the imino tautomeric form of TDP in catalysis is generally accepted (34–36). Given next are the experimental data indicating that in the resting state of the holotransketolase, the TDP aminopyrimidine ring exists in the imino tautomeric form and that this tautomer is the source of the inducible absorption band of holotransketolase. When TDP is placed into a medium less polar than water, a new band appears in its spectrum that was absent when the coenzyme was in aqueous solution (Fig. 5, curve 1). An analogous band in the near-UV region of the spectrum was detected when thiamine was dissolved in dioxane (37). It should be noted that in both cases the new absorption band appeared in the same region of the spectrum as the induced absorption band of holotransketolase (Fig. 2A). In the presence of phenylalanine, the intensity of the TDP absorption band in the ethanol solution is increased (Fig. 5, curve 2), consistent with the data obtained with mutant transketolase: the replacement of Phe445— located 4 A˚ away from the animopyrimidine ring of TDP (Fig. 1)—for isoleucine leads to a significant decrease in the intensity of the induced absorption band (33). Bearing in mind the hydrophobicity of the transketolase active site, the data on the induced absorption band of holotransketolase were interpreted as resulting from changed optical properties of the coenzyme following its incorporation into the hydrophobic pocket of the active center. Apparently the intensity of the induced absorption band increases at the cost of the interaction between the TDP aminopyrimidine ring and the Phe445 residue.


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Figure 5 Difference absorption spectra of TDP in ethanol solution (50%) in the absence (curve 1) and presence (curve 2) of phenylalanine with respect to TDP in water and TDP + phenylalanine in water, respectively. TDP, 2.5 mM; phenylalanine, 2.5 mM.

As is known, permanent interconversion of tautomeric forms of the TDP aminopyrimidine ring occurs during catalysis at the cost of the hydrogen bond formation between carboxyl group of Glu418 and the N1V atom of the TDP aminopyrimidine ring. Fast and easily reversible interconversion between these two forms of the TDP aminopyrimidine ring is a necessary requirement for the enzymatic catalysis (34,38). In aqueous solution, TDP occurs largely as aminoform (I in Scheme 2), having no absorption bands in the near-UV region (39). A hydrophobic environment promotes the formation of the iminoform (40) (II in Scheme 2). In the active site of holotransketolase, the aminopyrimidine ring of TDP is surrounded by hydrophobic amino acids (3,5). Therefore, we concluded that appearance of the new absorption band (Fig. 2A and B) results from transformation of the TDP to the imino form (II in Scheme 2) after incorporation of its aminopyrimidine ring into the hydrophobic pocket of the active site. In this case the amplitude of the new absorption band should depend on pH: a decrease in pH would shift the equilibrium from the imino form (II in Scheme

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Scheme 2 Tautomeric forms of TDP.

2) to the amino form (III in Scheme 2), with the consequent disappearance of the long-wave absorption. Indeed, as shown in Figure 6, the intensity of the induced long-wave absorption band of the TDP solution in ethanol decreases with an increase in the medium’s acidity. A similar pH dependence in the pH region 5.9–3.0 shows decreases in the amplitude of holotransketolase absorption (Fig. 7, solid curve) and CD band (Fig. 7, dashed curve): decreases with pH below 6.0. The only difference consists in the increase of the absorption in the pH range 7.6–5.9. The reason for this is currently unknown and needs further investigation. Notice the stability of holotransketolase in the investigated pH range and the absence of TDP release from the holoenzyme during the entire experimental time (data not shown). In the holoenzyme the imino form of TDP is stabilized not only by its interaction with Phe445, but also by its interaction with the glutamic acid residue through the N1V atom of the aminopyrimidine ring (36,40). In transketolase this residue is Glu418 (5,10). Indeed, the interaction of the transketolase mutant (where the Glu418 residue is replaced by glutamine or alanine) with TDP was accompanied by dramatic changes in the holoenzyme CD spectrum: Both positive and negative maxima were shifted toward shorter wavelengths (10). Together, the data presented here allow us to formulate a general concept on the nature of the induced optical band of holotransketolase and


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Figure 6 Dependence of the intensity of the TDP absorption band in the ethanol solution (50%) on the acidity of the medium. TDP, 2.5 mM.

the reason for its appearance. During the interaction of TDP with apotransketolase, the aminopyrimidine ring of the coenzyme incorporates into the hydrophobic pocket of the enzyme active site, after which its N4V-amino tautomeric form converts to the N1VH-imino tautomeric form, stabilized through the interaction with the Glu418 and Phe445 residues. The investigation of a variety of catalytically inactive TDP analogs has shown that either their interaction with transketolase is not accompanied by the appearance of the induced optical band or the character and intensity of optical changes differ from those occurring in the native holoenzyme (41). There is only one exception: N3V-pyridyl-TDP, in which the N1V atom was replaced by carbon. The CD spectrum of holotransketolase, bound to this analog, does not differ from those of the native holoenzyme (42). This is not surprising, since at certain conditions, the a-aminopyridine ring of this analog, like the aminopyrimidine ring of TDP, has the ability to convert into the imino form (43–45). The fact that TDP after the replacement of N1V for carbon (N3V-pyridyl-TDP) has lost its catalytic activity when incorporated into the holoenzyme, can be explained by the lack of the N1V atom, which is capable of forming a hydrogen bond with the Glu418 residue—a necessary requirement for catalysis.

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Figure 7 Effect of pH on the intensity of the holotransketolase absorption band, induced through TDP binding, on the absorption spectrum (solid curve) and CD spectrum (dashed curve). Glycyl-glycine buffer, 20 mM, pH 7.6; CaCl2, 2.5 mM; TDP, 2.5 mM; transketolase, 3 lM (solid curve) and 6 lM (dashed curve).

Generally recognized for all TDP enzymes is the immediate participation of the imino tautomeric form of TDP in the act of catalysis. It could be proposed, that in the case of transketolase, the coenzyme binds to the active site besides the V-conformation in the imino tautomeric form. It means that this form arises even at the step of the coenzyme’s binding to the apoprotein, prior to the catalysis. With other thiamine diphosphate enzymes, which do not possess this spectral properties, there may be another situation, and the imino form probably appears during catalysis only.

ACKNOWLEDGMENTS This work was partly supported by the Russian Foundation for Fundamental Research (grant No. 99-04-49121). We would like to thank Prof. G. Hu¨bner and Dr. R. Golbik for their collaboration in the stopped-flow experiments, Dr. L. S. Yaguzinsky, Dr. O. H. Spivey, and Dr. Aart de Kok for their helpful discussion.


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11 Structure of the A-Carbanion/ Enamine Reaction Intermediate in the Active Site of Transketolase, Determined by Kinetic Crystallography Tatyana Sandalova, Stina Thorell, and Gunter Schneider Karolinska Institutet, Stockholm, Sweden Erik Fiedler, Ralph Golbik, and Stephan Ko¨nig Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany

I. INTRODUCTION Transketolase is a ubiquitous TDP-dependent enzyme found in the nonoxidative branch of the pentose phosphate cycle. The enzymes from bacteria, yeast, and mammalian sources are homo-dimers with a molecular mass of about 74 kDa per subunit (1–3). The TDP molecule binds in a cleft at the interface between the two subunits and is, except for the C2 carbon atom of the thiazolium ring, completely inaccessible from the solution (4,5). TDP is bound in the V-conformation, a feature characteristic of all TDP-dependent enzymes (6,7). This conformation positions the 4V-amino group of the pyrimidine ring of TDP in close proximity to the C2 carbon atom of the thiazolium ring of the cofactor, an essential prerequisite for catalysis (4,6,8). 159

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Transketolase catalyzes ketol transfer between ketose and aldose sugars. The first half of the reaction cycle consists of the cleavage of the donor substrate and release of the first product, an aldose, and the formation of a covalently bound intermediate, the a-carbanion/enamine of a,h-dihydroxyethyl-TDP (DHETDP). The a-carbanion/enamine intermediate is of central importance in thiamine catalysis, because dependent on the nature of its substituent at the a carbon atom, a variety of enzymatic functions can be derived (9). Accordingly, the fate of this intermediate differs in the various TDPdependent enzymes; for instance, in pyruvate decarboxylase the a-carbanion will be protonated at the a-carbon position, leading to the expulsion of the product, acetaldehyde (8–10). In the transketolase reaction, the carbanion reacts with an acceptor substrate and the product, a ketose with the carbon chain extended by two carbon atoms, will be released (6,11). It is thought that the a-carbanion intermediate is stabilized via the thiazolium ring of TDP, which acts as an electron sink. A major source of stabilization is the neutral enamine, contributing to the resulting resonance hybrid (Fig. 1).

Figure 1 Formation of the a-carbanion/enamine intermediate using HPA as donor substrate.

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A peculiarity of transketolase is the acceptance of hydroxypyruvate (HPA) as donor substrate. Formation of the a-carbanion/enamine of DHETDP starting from HPA is in practice irreversible, because it is coupled to the decarboxylation of the substrate (Fig. 1). In the absence of a suitable acceptor substrate, the intermediate decomposes into enzyme-TDP and glycolaldehyde (12), albeit at a rather low rate. The crystal structure of the a-carbanion/enamine of DHETDP in the active site of transketolase could be determined at 1.9-A˚ resolution by trapping this intermediate using flashfreezing techniques (13). II. MATERIALS AND METHODS A. Protein Expression and Purification Recombinant TK was purified according to the protocol of Wikner et al. (14), with the modifications described in Ref. 12. B. Circular Dichroism Measurements The CD spectra of TK were recorded on an Aviv 62 CD spectrophotometer under conditions similar to the crystallization experiments (3.4 mg TK/mL in 100 mM glycyl-glycine, 1 mM TDP, 5 mM CaCl2, pH 7.9 and 40 mM HPA). The temperature was set to 4jC, and spectra were recorded at different incubation times using cuvettes with an optical path length of 1 cm. C. Kinetics of Formation of the TK-HPA Adduct The formation of the TK–intermediate complex formed through reaction with HPA was followed directly by the associated absorbance change at 300 nm (15) using a stopped-flow spectrophotometer at 25jC. D. Crystallography and Data Collection TK was crystallized by the vapor diffusion technique, as described previously (16), using PEG 6000 (13–17% w/v) as precipitant in the presence of 5 mM TDP and 5 mM CaCl2 (drop volume 15 AL). The reaction was started by incubating crystals in the cryo-protecting solution (50 mM glycyl-glycine buffer, containing 5 mM TDP, 5 mM CaCl2, 20% (w/v) PEG 6000, 20% (v/v) ethylene glycol, pH 7.9) at 4jC, including the donor substrate HPA (40 mM). The reaction was stopped by transfer of crystals into a nitrogen stream at 110 K at various time intervals. X-ray data were collected at beam line BW7B, EMBL outstation, DESY Hamburg, using crystals incubated for 30 s (2.37-A˚ resolution, Rsym=0.109) and 30 min (1.86-A˚ resolution, Rsym=0.052), respectively.

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E. Crystallographic Model Building and Refinement The model of holo-TK, refined to 2.0-A˚ resolution (5), was used as the source of initial phase information for the calculation of the electron density maps. Inspection of electron density maps and model building was carried out using the program O (17). Refinement was done with CNS (18). The refinement procedure started with simulated annealing, and the following cycles consisted of positional and B-factor refinement. Five percent of the diffraction data were set aside to monitor the progress by means of the free R-factor. Tight noncrystallographic symmetry restraints were imposed for all residues of the two polypeptide chains in the asymmetric unit. The protocol consisted of iterative rounds of refinement and model examination/rebuilding with O (17), until the Rfree value had converged. The final model contains amino acids 3 to 680 for each subunit of the dimer, two DHETDP molecules, two Ca2+ ions, and 854 (370 in the lower-resolution data set) water molecules. The final R values are 22.1% (30-s data set) and 19.8% (30-min data set), respectively. III. RESULTS AND DISCUSSION A. Circular Dichroism Measurement and Kinetics of HPA Binding to TK The binding of the cofactor TDP and the donor substrate HPA to TK can be monitored by near-ultraviolet CD spectroscopy (19,20). Holo-TK is characterized by a negative band in the CD spectrum at 320 nm. Inversion of this band is observed upon addition of donor substrate. The spectrum of holo-TK can be restored by addition of an acceptor substrate, indicating that the spectroscopic changes might be related to catalytic intermediates. This signal, proposed to correspond to the formation of the a-carbanion/enamine intermediate of (a,h-dihydroxyethyl)-TDP (19), was used as an analytical tool to follow the stability of the intermediate. As illustrated in Figure 2, the CD signal at 320 nm is stable in solution for at least 30 min under crystallization conditions. From the progress curves of the stopped-flow experiments, a rate constant of 44.6 mM1 s1 was calculated for the formation of the TKHPA complex and a rate constant of 0.0026 s1 for the release of glycolaldehyde. Decomposition of DHETDP is thus the rate-limiting step under these conditions. The kinetic and spectroscopic data thus indicate that in the absence of the acceptor substrate, the a-carbanion/enamine intermediate is relatively stable in the active site of transketolase. The use of the donor substrate HPA, coupled with an in-practice irreversible decarboxylation step, led to the accumulation of this intermediate, which allowed the determination of its threedimensional structure using cryocrystallography.


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Figure 2 Time dependence of the CD spectrum of holo-TK incubated with HPA (from top to bottom, after 30 s incubation, 9 min incubation, 15 min incubation, 30 min incubation, holo-TK without HPA).

B. Electron Density Maps and Overall Structure After reaction initiation with HPA, several data sets of presumptive TK– intermediate complexes were collected. The structures of these complexes were solved using difference Fourier methods and refined. The resulting electron density maps were of very good quality. The R-factors and the stereochemistry are as expected for models at the given resolution. The overall structure of the polypeptide chain in these complexes is very similar to the structure of holo-TK. Superposition of the 678 Ca atoms of the TK subunit results in an r.m.s. deviation of 0.3 A˚, and there are no local deviations larger than 0.5 A˚, even for side chains. Thus, there is no significant structural change in the structures described here that would have indicated a conformational transition upon formation of the intermediate.

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C. Structure of the DHETDP Intermediate The most significant feature in the initial Fo–Fc difference electron density maps calculated with the data sets collected 30 s and 30 min after reaction initiation is a strong, positive electron density extending from the C2 carbon atom of the thiazolium ring of TDP into the active site (Fig. 3), indicating a covalent adduct at this carbon atom. A model of the a,h-dihydroxyethyl moiety of DHETDP could be fitted straightforwardly into this difference electron density, and the resulting maps after refinement clearly indicate successful trapping of the a-carbanion/enamine intermediate in the active site of transketolase (Fig. 3). After refinement, the B-factors for the atoms of the a,hdihydroxyethyl moiety (22.8-A˚2) are very similar to those for TDP (19.9-A˚2) and surrounding active-site residues, indicating stoichiometric formation of the intermediate. The results are very similar for the data sets collected 30 s and 30 min after reaction initiation. The crystallographic data show that the atoms of the thiazolium ring, as well as the Ca, Ch, and the Ca oxygen atoms of the dihydroxyethyl moiety,

Figure 3 Refined 2Fo–Fc electron density map for the bound a-carbanion/enamine of DHETDP in the active site of transketolase, contoured at 1.0j, for the data set collected 30 min after reaction initiation. The refined model is superimposed.


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are all in one plane. The electron density fits best to a planar structure of DHETDP, i.e. sp2 hybridization of the a-carbon atom, suggesting a predominant enamine character of the intermediate (Fig. 1). Attempts to model the intermediate with sp3 geometry at the a-carbon atom are not consistent with the electron density and also result in a small, but significant, increase in the free R-factor by 0.4%. The carbanion/enamine has the E-configuration, as predicted from molecular modeling of decarboxylation of pyruvate in pyruvate decarboxylase (21). Furthermore, the formation of the covalent intermediate does not result in changes in conformation of the cofactor itself, compared to the TDP conformation observed in the structure of holotransketolase. The a-hydroxyl oxygen is close (2.9 A˚) to the 4V-NH2 group of the aminopyrimidine ring of TDP, albeit at an angle (74j between the Ca carbon, the oxygen at the Ca, and the 4-amino nitrogen) unfavorable to the formation of a strong hydrogen bond. Such a hydrogen bond has been proposed as a key interaction of central importance to catalysis. The essential function of the 4VNH2 group had been demonstrated by showing that deamino-TDP is catalytically inactive (22). Although in the structure of the a-carbanion/enamine intermediate the geometry is not favorable for a hydrogen bond, this situation will be different in the steps immediately preceding and subsequent to the formation of DHETDP. During the initial nucleophilic attack of the C2 carbanion on the carbonyl carbon of the donor substrate, a negative charge will develop at the carbonyl oxygen atom and the hybridization at the carbonyl carbon atom will change from sp2 to sp3. This leads to a geometry in the transition state and in the first covalent adduct that is more favorable for hydrogen bond formation between the a-hydroxyl oxygen and the 4V-NH2 group. At this stage the 4V-NH2 group can act in proton transfer to this oxygen atom in a similar manner as described for the deprotonation of the C2 carbon of the thiazolium ring of TDP (23). In this way, the negative charge of the oxygen atom at the a-carbon is compensated. The structure of the a-carban-carbanion/enamine intermediate described here thus supports mechanistic proposals pinpointing the importance of the 4V-NH2 group of TDP in proton transfer steps during catalysis (6,8,22). D. Interactions of DHETDP with Active-Site Residues None of the side chains interacting with the a,h-dihydroxyethyl moiety of DHETDP changes its position compared to the structure of holo-TK, and they are thus already poised, in the absence of the donor substrate, to interact with the a-carbanion intermediate. The interactions of the TDP moiety with residues in the cofactor-binding site also remains unchanged. The a,h-dihydroxyethyl moiety is held in place through a number of hydrogen bonds made to surrounding amino acids (Fig. 4). The h-hydroxyl

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Figure 4 Surroundings of the reaction intermediate DHETDP in the active site of transketolase. Hydrogen bonds (distances 600 s1 (45), and in activated E477Q and D28A variants it is 30 s1 at 5jC, fast enough not to be rate limiting in any of the three cases. D. Additional Aspects of the Alternating-Sites Mechanism We suggest that the active sites in the functional dimer are not acting independent of one another, and the pre- and postdecarboxylation phases of the reaction are tightly coupled. With this coupling, the alternating-sites model could explain: (a) equal rates of pre- and postdecarboxylation in wildtype YPDC, since two parts of the reaction are synchronized; (b) apparent participation of all four acid–base groups in both pre- and postdecarboxylation steps, as suggested by the steady-state kinetics reported (44), according to which both V/K and V-type kinetic terms are affected by substitutions at D28, H114, H115, and E477. E. Multiple Regulatory Mechanisms in TDP-Dependent Decarboxylases It is useful to place these proposals in perspective concerning the several different regulatory mechanisms already delineated for TDP-dependent enzymes in general and 2-oxo acid decarboxylases in particular. The regulation of the mammalian 2-oxo acid dehydrogenases by a kinase/phosphatase system is discussed elsewhere in this volume but does not apply to the simpler decarboxylases. Several enzymes were shown to possess hysteretic binding of the Mg(II) and, bonded to it by inner-sphere complexation, an oxygen from both the aand h-phosphate of the diphosphate side chain of TDP. This could be clearly demonstrated on the E1 component of the E. coli pyruvate dehydrogenase multienzyme complex (46). All of the YPDCs (but not ZmPDC) are subject to substrate activation. The current state of our understanding was discussed earlier. The signal is triggered by the substrate’s being bound to C221 and is likely propagated via H92 to E91 to W412, to the residue adjacent to G413, the latter forming a strong hydrogen bond to the 4V-amino group of TDP. This could be construed as an ‘‘intrasubunit’’ pathway. Studies in our laboratory on the E1 compo-

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nent of the E. coli pyruvate dehydrogenase multienzyme complex (PDHc-E1) also pointed to a substrate activation phenomenon (47), whose structural origins are still unknown. The results on YPDC have led us to conclude that there is yet an additional signal transduction pathway, in which there is an interaction of active sites and in which the residue D28 from one subunit and residue E477 of the second are major players in a functional dimer. Complementation of the very low-activity D28N and E477Q variants to virtually fully active species provides some support for this notion as well (48). We conceive of this novel signal transduction pathway as an ‘‘intersubunit’’ pathway, to distinguish it from the substrate activation model. Our recent studies on BFD also led us to a similar proposal on the basis of pre-steady-state kinetics (40,41). We recently also obtained evidence for such an alternating active-site mechanism on YPDC from pre-steady-state kinetic data, with a chromophoric substrate (pyridine-based) enabling detection of reaction time courses for both LTDP and enamine-like intermediates (49). Elucidation of further details of this novel pathway, whether the ‘‘intrasubunit’’ pathway and ‘‘intersubunit’’ pathway are interconnected, and, of course, as to how general this ‘‘intersubunit’’ pathway is for TDP enzymes, remain challenges for the future. V. EVIDENCE FOR THE PRESENCE OF THE 1VV, 4VV-IMINOTDP TAUTOMER ON YPDC As discussed in Section II, one of the properties of the V TDP conformation is to bring to within 3.4 A˚ the N4V and C2 atoms. This observation tempted others and us to suggest that there is intramolecular proton transfer between these two atoms (Scheme 1) and that the 4V-aminopyrimidine ring cycles between the 4V-amino and 1V,4V-imino tautomeric forms during the reaction sequence, in a manner suggested 25 years ago (14,50,51). This rare imino tautomer is stabilized by three highly conserved hydrogen bonds to N1V, N3V, and N4VH3V (denoting the proton bonded to N4V on the N3V side of the pyrimidine ring). In 2002, the Rutgers group reported both rapid-scan stopped-flow UV and circular dichroism (CD) evidence for a hitherto-unreported absorption between 300 and 310 nm on the E477Q YPDC variant in the presence and even in the absence of the pyruvate substrate (15). A model for the absorption was generated by adding a base to N1-methylpyrimidinium salts in either water or aprotic organic solvents. On the basis of this model system, it was suggested that the absorption on YPDC between 300 and 310 nm pertain to the 1V,4V-iminopyrimidine tautomer of TDP (15). Concurrently, a group at

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Figure 11 Circular dichroism spectra of the E477Q YPDC variant in the presence of acetaldehyde in the absence and presence of pyruvamide. Enzyme was dissolved to a concentration 8.6 mg/mL (or 143 AM active sites). Acetaldehyde was added to 0.3 M concentration at 25jC. Pyruvamide was added to a final concentration of 20 mM. Upper panel represents actual spectra; difference spectra are shown in the lower panel.


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Moscow State University suggested that the broad negative CD signature centered around 320–330 nm, and observed for three decades on TK and on mammalian PDHc’s, is indeed pertinent to the 1V,4V-imino TDP tautomer (52). Importantly, we have identified conditions under which this negative CD signature centered at 320–330 nm could be observed on YPDC (Fig. 11) and on PDHc-E1 from E. coli (to be published) for the first time. However, these observations typically required so-called ‘‘ligation’’ conditions, i.e., the presence of an acceptor for the enamine. This raises the interesting further issue, should this negative CD band at 320–330 nm indeed be associated with the 1V,4V-imino TDP tautomer, of why and how the protein stabilizes it under these particular conditions. In recent work, the positive CD band has also been observed on PDHcE1 in the presence of the phosphonolactyl-TDP, a stable analog of LTDP, strongly suggesting that with this analog the TDP exists in its imino tautomeric form (see Chapter 23 in this volume). At the same time, we have also shown that the positive CD band centered at 305–310 nm and the negative one near 320–330 nm exist under different conditions. While we are confident of the assignment of the positive band at 305–310 nm to the fixed V-conformer of the 1V,4V-iminoTDP, we are less certain of the origins of the negative band at 320–330 nm. It is clear, however, that both bands are associated with bound TDP since there is little else conserved in the active centers of the four enzymes in which they have been observed (TK, PDHc from both mammalian and bacterial sources, YPDC). The only pertinent kinetic observation of the 1V,4V-iminoTDP so far has been on the E477Q YPDC variant, as derived from pyruvate. Now that we have identified a spectroscopic signature both in the UV and in the CD spectrum, we consider that the existence of this tautomer is now experimentally established and that any complete TDP mechanism needs to involve the species.

VI. ASSIGNMENT OF FUNCTION TO ACTIVE-CENTER RESIDUES: METHODS AND RESULTS A. pH Dependence of Kinetic Constants The four active-center groups with potential acid–base properties in the region of pH optimum of YPDC have been studied with the substitutions D28A, H114F, H115F, and E477Q introduced by site-directed mutagenesis methods. The steady-state kinetic constants were determined in the pH range of activity for the enzyme (44). The substitutions result in large changes in kcat and kcat/S0.5 (and related terms), indicating that all four groups have a role in transition-state stabilization. Furthermore, these results also imply that all

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four are involved in some manner in stabilizing the rate-limiting transition state(s) both at low substrate (steps starting with substrate binding and culminating in decarboxylation) and at high substrate concentration (steps culminating in product release). With the exception of some modest effects, the shapes of neither the bell-shaped kcat/S0.5–pH (and related functions) plots nor of the kcat–pH plots are changed by the substitutions. Yet the fractional activity remaining after substitutions virtually rules out any of the four residues as being directly responsible for initiating the catalytic process by ionizing the C2-H. There is no effect in C2H/D exchange rate exhibited by the D28A and E477Q substitutions. These results strongly imply that the base-induced deprotonation at C2 is carried out with the participation of the only remaining base, the 1V,4V-iminoTDP tautomer of the coenzyme. B. Carboligase Side Reactions YPDC, in addition to forming its metabolic product acetaldehyde, can carry out carboligase reactions in which the central enamine intermediate reacts with acetaldehyde or pyruvate (instead of the usual proton electrophile), resulting in the formation of acetoin and acetolactate, respectively (typically, 1% of the total reaction). Due to the common mechanism shared by the acetaldehyde-forming and carboligase reactions through decarboxylation, a detailed analysis of the rates and stereochemistry of the carboligase products formed by the E477Q, D28A, and D28N active-center YPDC variants was undertaken. While substitution at either position led to an approximately two to three orders of magnitude lower catalytic efficiency in acetaldehyde formation, the rate of acetoin formation by the E477Q and D28N variants was higher than that by wild-type enzyme (41). Comparison of the steadystate data for acetaldehyde and acetoin formation revealed that the ratelimiting step for acetaldehyde formation by the D28A, H114F, H115F, and E477Q variants is a step postdecarboxylation. In contrast to the wild-type YPDC and the E477Q variant, the D28A and D28N variants could synthesize acetolactate as a major product, with an activity higher than that reported for the enzyme acetolactate synthase in plants and bacteria. The lower overall rate of side-product formation by the D28A variant than wild-type enzyme attests to participation of D28 in steps leading up to and including decarboxylation. The results also provide insight to the state of ionization of the side chains examined (discussed earlier in this review). The two YPDC variants could also be used for enzyme-catalyzed synthesis of a-ketols. (R)Acetoin is produced by the E477Q variant with greater enantiomeric excess than by wild-type YPDC. (S)-Acetolactate is the predominant enantiomer produced by the D28-substituted variants, the same configuration as produced by the related plant acetolactate synthase (41).


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C. Intermediate Partitioning Experiments 1. Lactyl-TDP The first TDP-bound intermediate in Scheme 1 is the result of a nucleophilic attack at the carbonyl carbon of the substrate and produces LTDP. Formation of this intermediate is consistent with the driving force for decarboxylation being the electrophilic (electron-withdrawing) effect of the thiazolium ring beta to the departing carboxylate. This intermediate is highly reactive (especially in its zwitterionic form), and its chemistry was well established by Kluger and coworkers, who also reported that pyruvate decarboxylase failed to decarboxylate it (53). Two methods have recently been reported that indeed indicate that this intermediate is distinct. Tittmann and Hu¨bner and their group of (54; see also Chapter 5 in this volume) carried out acid quench of a mixture of pyruvate, TDP, and YPDC, showing that one can observe both LDP and HETDP under these conditions, strongly suggesting that both are on the pathway. At Rutgers, a different approach was developed. First, it was shown that the E91D YPDC variant could form a stable apo enzyme, which could be reconstituted with virtually any TDP derivative, including LTDP (29). Next, it was found that LTDP is partitioned on apo-YPDC (Scheme 5), both to pyruvate (in the reverse direction), and is indeed decarboxylated

Scheme 5 Intermediate partitioning experiments on YPDC.

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according to the measurement of enamine and acetaldehyde. This evidence strongly supports the intermediacy of LTDP as a distinct chemical entity on pyruvate decarboxylases (55,56). 2. HETDP This intermediate is in acid–base equilibrium with the enamine intermediate, the central intermediate on all TDP-dependent catalytic pathways. The Rutgers group has spent the past 20 years elucidating the properties of this enamine intermediate in the absence and in the presence of TDP enzymes (4,5). These are very important comparisons if we are to understand the contribution of the protein to individual steps. By direct observation of the enamine generated in a stopped-flow spectrophotometer, the rate constants for reversible proton dissociation at the C2a position could be measured in water. The pKa is between 15–16 for C2a-hydroxybenzylthiazolium salt (57,58), and near 15 in 32–37 mole % DMSO for C2a-hydroxyethylthiazolium salt (extrapolates to approximately 18 in water; Ref. 59). It was concluded that YPDC and BFD assist in the protonation of the enamine to afford rate constants commensurate with enzymatic turnover numbers. In view of the difficulty in generating the enamine in aqueous solution, we became interested in whether the enzymes could overcome this high pKa problem. The partitioning experiment is outlined in Scheme 5. When the E91D variant of apo-YPDC was exposed to C2a-hydroxybenzylTDP (HBTDP), this putative intermediate was partitioned on the enzyme between release of the benzaldehyde product (evidenced by regeneration of active enzyme) and dissociation of the proton at C2a to form the enamine/ C2a-carbanion intermediate (evidenced by the appearance of the visible spectrum of the intermediate). While the pKa for this dissociation is f15.4 in water, formation of the enamine at pH 6.0 on YPDC indicates a greater than 9-unit pKa suppression by the enzyme environment (60). Using the fluorescence emission properties of thiochrome diphosphate, a fluorescent TDP analog and a competitive inhibitor for YPDC, an apparent dielectric constant of 13–15 was estimated for the YPDC active center (calibrated against a series of 1-alkanols). Such a low effective dielectric constant could account for much of the observed >9-unit pKa suppression at the C2a position for ionization of HBTDP. The dramatic stabilization of this (and presumably other) zwitterionic/dipolar intermediate(s) is sufficient to account for as much as a 109-fold rate acceleration on YPDC, providing a significant contribution to the rate acceleration by the protein over and above that afforded by the coenzyme. Similar experiments have also been carried out with HETDP (56), confirming the ability of YPDC to partition this intermediate as well, to acetaldehyde in the forward and the enamine in


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the reverse direction (the enamine in this case has a Emax near 295 nm, so it could be detected only by indirect oxidative methods). With this intermediate, whose pKa is even higher, perhaps 18, the pKa suppression induced by the enzyme is even more impressive. At the same time, the enamine could also be generated from HETDP by PDHc-E1. This result suggests that the PDHc-E1 also possesses an active center that can stabilize zwitterions/ dipolar ions.

D. Specific Examples of Functional Assignments 1. E51, a Conserved YPDC Residue Hydrogen Bonded to the N1V TDP Atom On the basis of the X-ray structure alone, one would surmise that this highly conserved residue will have a dramatic effect in all TDP enzymes, since there is a good hydrogen bond formed between the N1V atom and the carboxylate oxygen atom. Yet at the resolution of the X-ray structures currently available for TDP enzymes, the hydrogen positions are not defined, so neither the state of ionization nor the tautomeric state of TDP could be deduced with any certainty. This is generally true for all acid–base residues, so modeling with the assumption of a particular state of ionization or of the tautomeric state would be futile, given the uncertainty of the effects of the microenvironment on the pKa’s. On YPDC, the E51Q, E51D, E51N, and E51A substitutions all led to greatly diminished kcat and kcat/Km values (18). The E51D substitution turned out to be informative, since at low substrate concentrations (steps starting with free enzyme and culminating in decarboxylation), the log kcat/S0.5–pH plots displayed an acid shift for the entire curve, showing that the distance of the negative charge from N1V influences this curve. A plausible explanation is that the pKa of the N1V-protonated pyrimidinium ring is reduced (i.e., the ring is more difficult to protonate), this in turn making it more difficult to catalyze the tautomeric equilibration, which, in turn would reduce the rate at which the 1V,4V-imino TDP can abstract the thiazolium C2H to form the ylide/carbanion in the required first step. It had been reported that the E51Q substitution reduced the rate of C2H-to-D exchange (as a measure of the rate of the first-step ylide formation) significantly, but even for that variant C2H dissociation may not have become rate limiting (45). A different type of experiment was designed to test the E51A variant (low but detectable activity) with TDP and N1V-methylTDP in parallel. The premise of the experiment is shown in Scheme 6, testing whether the activity requires both tautomers and the N1V-protonated pyrimidinium intermediate, an obligatory species for the interconversion of the two tautomers. The N1V-

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Scheme 6

methylTDP is an electrostatic mimic for the N1V-protonated intermediate, and it can be converted to the imino tautomer, as we have shown, but it precludes existence of the 4V-aminopyrimidine form. Addition of N1V-methylTDP to the wild-type YPDC or the E51A variant led to no observable activity, but it was found that N1V-methylTDP bound to Apo-E51A some 18 times better than did TDP itself. We conclude from this that all three forms, including the protonated TDP and both tautomers, must coexist on YPDC, not possible for the N1V-methylTDP, which in fact is bound in a satisfactory manner. The conserved residue is needed to catalyze the tautomeric equilibration, but in its absence the tautomerization is still possible, because all of the variants at this position displayed residual activity. Assuming flexibility


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of the enzyme in the region of E51, in theory a water molecule could also catalyze the tautomeric equilibration, albeit at a slower rate. In accord with this idea, incubation of the E51A (but not of the other E51-substituted variants) with TDP overnight increased the activity significantly. 2. D28, a Residue Conserved Only in Pyruvate Decarboxylases In the structures of the pyruvate decarboxylases from yeast (1,2) and Zymomonas mobilis (61), this residue is located above the 4V-aminopyrimidine ring with a water molecule nearby. While both kcat and kcat/Km-type terms are significantly reduced for the YPDC D28A and D28N variants, its clear participation in a step postdecarboxylation became evident from studies of the carboligase side reaction. Remarkably, though these substitutions led to greatly diminished acetaldehyde production, they had no adverse effect on the carboligase side reactions; in fact, this was the only substitution identified so far that converted the enzyme to a fairly respectable acetolactate synthase (42). From these observations, we deduced that the state of ionization is D28COOH through formation of LTDP, but D28 transfers a proton thereafter to E477, thus becoming D28COO, so as to repel the second pyruvate in the wild-type enzyme. This would account for the fact that with wild-type enzyme the carboligase side products constitute less than 1% of the total product. The strongest evidence for a postdecarboxylation role was generated from studies in which we partitioned HETDP on the apo-E91D YPDC variant (Scheme 5, bottom): Enamine p HETDP ! Acetaldehyde We then tested partitioning of HETDP with the doubly substituted D28A/ E91D variant and found that this substitution only (among D28, E477, H114, and H115) allowed virtually no formation of acetaldehyde (55,56). We concluded that a major role of residue D28 is to help deprotonate the C2a-OH for release of acetaldehyde. The residue D28 on YPDC also appears to have a role in the protonation of the enamine, perhaps in conjunction with residue E477, according to partial reduction in the rate of enamine formation from HETDP by the D28A/E91D variant. Consistent with this suggestion, in careful difference CD spectra (Fig. 12), the D28A and E477Q variants appear to give rise to some of the same and one different long-lived chiral intermediates, both different from that observed with the wild-type YPDC (15). The positive CD signal of this intermediate was centered at 290 nm, very similar to the wavelength determined for the enamine derived from pyruvic acid (295 nm) in models, consistent with the idea that this residue helps to protonate the enamine.

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Figure 12 CD difference spectra of YPDC-bound intermediates on wild-type and variant enzymes: on the left, protein +/ substrate; on the right, the spectra on the left are subtracted from each other.

3. E477, a Residue Located Over the Thiazolium Ring of Pyruvate Decarboxylases The E477Q substitution on YPDC led to a greatly diminished acetaldehyde release with very much reduced kcat and kcat/Km (44). However, not only was the carboligase reaction leading to acetoin not impaired, but, to the contrary, its rate was even faster than with the wild-type enzyme. In essence, the E477Q substitution converted this variant to an acetoin synthase (42). This behavior suggests a role subsequent to the decarboxylation step (enamine formation), since, through decarboxylation, acetaldehyde and acetoin formation share a common pathway. More insight to the behavior of this residue was gleaned from rapid-scan stopped-flow and difference CD spectroscopic measurements, indicating the buildup of an intermediate with Emax near 310 nm and positive CD signal centered at 305, respectively (15,42). These observations suggest the presence of the 1V,4V-iminoTDP tautomer (see earlier). According to the intermediate partitioning experiments mentioned earlier, addition of HETDP to the apoE91D/E477Q variant allowed normal release of acetaldehyde but an impaired rate of enamine formation. This evidence suggests that E477 at least contributes to the enamine protonation step. As suggested by Figure 12, there is also enamine buildup according to the CD experiment with the E477Q YPDC variant. For this variant, there is a difference peak somewhat broader than with the D28A variant and at a slightly longer wavelength, since it likely represents a superposition of a signal for the enamine at 295 nm and the one for the 1V,4V-iminoTDP at 305 nm. On the basis of the electronic spectroscopic data indicating that with the E477Q YPDC variant the lifetime of the 1V,4V-iminoTDP is increased, it


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appears that the E477 may assist proton transfers to and from the imino TDP, perhaps deprotonating the 4V-amino group, possibly with the intermediacy of a water molecule. This could be one of the most important and unique functions of residue E477, and it may explain why the steady-state kinetic data indicate the participation of E477, along with the imino-TDP, in reactions starting with free enzyme through decarboxylation and those culminating in product release. VII. SUMMARY From these studies on YPDC, a number of points of general significance to TDP enzymes have emerged. According to both X-ray studies and solution kinetic studies, there may be multiple conformations of the enzyme; certainly in solution there appears to be more than one active species. It is also likely that the energy barriers interconverting the various conformations/tetramer assemblies are small. There is accumulating evidence that both YPDC and BFD behave according to an alternation of active sites in a functional dimer mechanism. This raises the issue of whether other ThDP enzymes also follow this mechanism. The role of the protein in YPDC catalysis is also becoming clearer. Yet, as seen earlier, assignment of a role to a particular amino acid residue in catalysis at individual steps of the pathway is challenging, while assignment of the state of ionization to an acid–base residue in individual steps is daunting. The only residue where we have achieved this is D28, which undergoes at least one change in ionization state on the pathway. This should serve as a warning to those drawing conclusions about such issues by inspection only. With the recent observation of the 1V,4V-imino TDP tautomer in both YPDC and PDHc-E1, we have an additional intermediate on the reaction pathways that must be taken into account. The role of the protein is clearly to stabilize this and other intermediates, and the associated transition states. It is worth emphasizing that starting with the ylide, progressing to LTDP, to the enamine, to the C2-a-oxyethylTDP (the presumed alkoxide intermediate for acetaldehyde release from HETDP), all of these intermediates carry a positive and a negative charge, only partial on the ylide and the enamine, but full at the other two. We believe that the evidence from our laboratory in which the enamine could be generated from HETDP on both YPDC and PDHc-E1 is strong in favor of an ‘‘environmental’’ effect that produces the observed pKa suppressions. While several residues may participate in proton transfer to and from the C2a atom of HETDP on YPDC, no such steps exist on the pathway of PDHc-E1; hence, with the latter the observations must be due to such an environmental effect. For YPDC, we designed an experiment to enable estimation of the effective dielectric

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constant where the value would be sufficient to account for the observed pKa suppression. Finally, we continue to be astonished by the catalytic versatility displayed by TDP: (1) electrophilic catalysis, also mimicked in solution studies using the thiazolium salts by themselves, and (2) intramolecular acid–base catalysis via the amino-imino tautomerization, enforced and afforded by the protein environment. So far, thiamine diphosphate is unique among coenzymes in utilizing such a dual catalytic apparatus.

ACKNOWLEDGMENTS Supported at Rutgers by NIH-GM-50380 and the NSF Training Grant -BIR 94/13198 in Cellular and Molecular Biodynamics (FJ, PI), at Pittsburgh by NIH-GM-61791 and VA Merit Review (to WF, PI).

ABBREVIATIONS USED TDP YPDC WT YPDC TK HETDP LTDP CD PDHc ZmPDC BFD E. coli PDHc-E1 HBTDP POX

thiamin diphosphate yeast pyruvate decarboxylase wild-type YPDC transketolase C2-a-hydroxyethylthiamin diphosphate 2-a-lactylthiamin diphosphate circular dichroism pyruvate dehydrogenase multienzyme complex pyruvate decarboxylase from Zymomonas mobilis benzoylformate decarboxylase Escherichia coli pyruvate dehydrogenase, the first subunit of PDHc (TDP dependent) C2-a-hydroxybenzylthiamin diphosphate pyruvate oxidase

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13 Solvent and Carbon Kinetic Isotope Effects on Active-Site and Regulatory-Site Variants of Yeast Pyruvate Decarboxylase Wen Wei, Min Liu, Lan Chen, W. Phillip Huskey, and Frank Jordan Rutgers University, Newark, New Jersey, U.S.A.

I. INTRODUCTION The reaction catalyzed by yeast pyruvate decarboxylase (YPDC) contains three fundamental processes, as shown in Scheme 1 (1–4). The coenzyme thiamine diphosphate (TDP), which is tightly bound to the enzyme, first adds to the substrate pyruvate to form the intermediate known as C2a-lactylthiamin (LTDP). This intermediate is poised for the second process, decarboxylation, with the thiazolium ring serving as an electron sink. The decarboxylation produces a second key intermediate, the enamine, which undergoes protonation at C2a, followed by elimination of acetaldehyde and regeneratation of the coenzyme. YPDC is subject to regulation by its own substrate, as is indicated in the steady state by sigmoidal plots of steady-state velocity vs. pyruvate concentration. The regulation is thought to be triggered by pyruvate binding at a regulatory site some 20 A˚ away from the active site, where a second molecule of pyruvate is decarboxylated (5–7). Crystal structures of the enzyme, a tetramer with four active sites, have been published (8). 217

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Scheme 1 Chemical steps in the pyruvate decarboxylase reaction. Product-release steps are shown as irreversible steps in accordance with the initial velocity and coupledenzyme assay conditions of the experiments.

We have used kinetic solvent isotope effects and kinetic carbon isotope effects on reactions catalyzed by wild-type YPDC and active-site and regulatory-site variants to learn about the role of individual amino acid residues in catalysis and regulation. These studies complement recent work directed at the same problem using rapid-quench techniques to determine the steadystate concentrations of the key reaction intermediates in catalysis by pyruvate decarboxylases from Zymomonas mobilis and yeast (9). A. Transition-State Theory of Kinetic Isotope Effects The analysis of kinetic isotope effects on enzymatic reactions usually begins with a treatment based on transition-state theory. Equation (1) summarizes this treatment in terms of isotopic fractionation factors (f) for the reactant state (RS) and transition state (TS) corresponding to the rate constant (k). The isotopic fractionation factors are equilibrium constants for isotopic ex-

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change reactions, defined to reflect the preference for the heavy isotope over a light isotope at a specific molecular site relative to a reference site. When the same reference site is chosen for the reactant state and transition state, Eq. (1) shows that the kinetic isotope effect is a relative measure of the affinity of isotopes in the two states. The origin of the isotopic selectivity of a particular molecular site lies in the isotopic differences in the translational, rotational, and vibrational energies, and in many cases informative qualitative analyses can be formulated based only on ideas about a few vibrational modes. For the carbon isotope effects and solvent isotope effects of this work, we use this standard transition-state approach. In cases that involve rate-limiting transfers of a proton, hydride ion, or hydrogen atom, quantum mechanical tunneling can be so important that a simple transition-state analysis may not be valid (10,11). nRS Y

KIE ¼ kðlightÞ=kðheavyÞ ¼

i¼1 nTS Y

/i ð1Þ /j

j¼1

/x ¼

½X heavy=½X light ½Reference heavy=½Reference light

ð2Þ

Some useful isotopic fractionation factors for solvent isotope effect studies are / f0.5 for the thiol SH and / f0.2–0.7 (25jC) for hydrogen bonds that are stronger than those of the reference, bulk water (12,13). Using these fractionation factors and Eq. (1), kinetic solvent isotope effects can be readily estimated for several cases. If, for example, a reactant-state thiol proton is lost to the bulk solvent in the transition state, the observed solvent isotope effect should be inverse (kHOH/kDOD 1) solvent isotope effect is the formation of a strong hydrogen bond in a rate-limiting transition state. If / = 0.33 for one of these transition-state hydrogen bonds, the observed solvent isotope effect would be 1/0.33 = 3.0, according to Eq. (1). Hydrogen bonds of this type could describe the proton transfers in transition states for general acid or base catalysis (14–16). For carbon isotope effects on decarboxylation reactions, /RS//TS f1.05 for many rate-limiting C–C cleavage reactions provides a useful benchmark for k12/k13 measurements. When decarboxylation limits a reaction rate, carbon kinetic isotope effects are large at about 1.05 because the C–C stretching vibration, a source of isotopic selectivity in the reactant state, is missing in the transition state (17).

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B. Rate-Limiting Steps in the Wild-Type Yeast Pyruvate Decarboxylase Reaction The rate-limiting steps for wild-type YPDC have been established using kinetic isotope effects (18,19) and reviewed (2). The minimal mechanism of Alvarez, Ermer, Hu¨bner, Schellenberger, and Schowen (18), shown in Scheme 2, accounts for the steady-state kinetics of wild-type YPDC at its pH optimum of 6.0 with four steady-state parameters, kcat, kcat/A, kcat/B, and kcatKi, displayed in inverse form on the scheme. The relative importance of selected pairs of reactant and transition states for each of these kinetic parameters is shown, in the bottom section of Scheme 2. The third-order rate constant, kcat/ A, is shown, for example, as being determined by the difference in the free energy at the state of the free enzyme and a transition state corresponding to a step involved in formation of the LTDP intermediate. As can be seen by inspection of the chart for the remaining steady-state parameters, the decarboxylation transition state contributes to kcat/B, in a minor way, and it is one of two equally important transition states for kcat. C. Steady-State Kinetics of Active-Site and Regulatory-Site YPDC Variants The three active-site YPDC variants described here are modifications of wildtype carboxyl functions: E477Q (glutamic acid to glutamine), D28A (aspartic acid to alanine), and E51D (glutamic acid to aspartic acid). As is shown in Figure 1, E477 and D28 are near the C2 carbon of the thiazolium ring and located near where pyruvate is likely to bind in the active site. E51 is located within hydrogen-bonding distance from a nitrogen atom (N1V) of the TDP 4Vaminopyrimidine ring. Ionized E51 has been proposed to activate the imino tautomer (20–24) of the pyrimidine ring to serve as a base in removing the C2H proton of the thiazolium ring (25). All three of these active-site variants show steady-state kinetics that are qualitatively similar to the wild-type YPDC, although a quantitative accounting for substrate inhibition and carboligase side reactions requires a more complex mechanistic model (26).

Scheme 2 A minimal mechanism and rate-limiting steps for the wild-type enzyme as established by Schowen and coworkers (Refs. 2,40). The fractional contribution of processes identified by pairs of reactant states and transition states is shown in the table for each steady-state rate parameter. A value of 1.00 indicates that a given process is entirely rate limiting; a value of 0.00 would indicate that the process is not involved in determining a given rate parameter. (Steady-State Kinetics: mechanism of Alavarez, et al. [Ref. 18]; Rate-Limiting Steps: based on rate constants in Schowen [Ref. 2].)


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Figure 1 Carboxyl side chains in the yeast pyruvate decarboxylase active site, drawn using Molscript (Ref. 42) and crystal-structure coordinates (Ref. 8).

The dependence of the steady-state velocity on pyruvate concentration is sigmoidal in pyruvate, and substrate inhibition is prominent. The turnover numbers for these active-site variants are 300–600 times smaller than the value for the wild-type enzyme. Modification of a cysteine residue (C221), long thought to be important in the regulation of yeast YPDC, produces significant changes in the steadystate kinetics that are consistent with assignment of this amino acid as the locus of the regulatory site (27,28). The sigmoidal part of the substrate concentration dependence of the steady-state velocity, seen for the wild-type enzyme, is not seen for any of the C221 variants studied to date. These are all double mutants in which the identity of a second nearby cysteine (C222) is changed to alanine (A) to eliminate the possibility for functional compensation when C221 is modified. The three regulatory variants described here are identified as C221A/C222A, C221D/C222A, and C222E/C222A, corresponding to substitutions of cysteine-221 for alanine (A), aspartic acid (D), and glutamic acid (E). The turnover numbers for these regulatory variants are about half the value for wild-type YPDC.


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II. RESULTS A. Solvent Isotope Effects To obtain the solvent isotope effects shown in Table 1, steady-state parameters were first determined from least-squares fits of steady-state velocities vs. pyruvate concentration. For the wild-type YPDC and for the active-site variants, two types of fitting equations were used. One corresponds directly to the model of Scheme 2 and gives kcat, kcat/A, kcat/B, and kcatKi as steady-state parameters. The other fitting equation used was a Hill-type (29) equation [Eq. (3)] modified to account for substrate inhibition: v0 =½E0 ¼ kcat =ðSn0:5 þ ½Sn ð1 þ ½S=Ki ÞÞ

ð3Þ

The key parameters for these fits are kcat, kcat/S0.5 (a constant with secondorder units corresponding to the rate at half the maximal rate), and the Hill coefficient, n. The steady-state parameters for both types of fits were determined for a series of pH and pD values in H2O and D2O buffers, and these data sets were fit to simple models for bell-shaped pH rate profiles to determine the pH- or pD-independent rate parameters. The ratios of these parameters are shown as the solvent isotope effects in Table 1. The regulatory-site

Table 1 Solvent Kinetic Isotope Effects on Wild-Type and Site-Specific Variants of Yeast Pyruvate Decarboxylasea DOD

(kcat)

WTc(28)

1.46 F 0.08

C221Ad (39) C221Dd (28) C221Ed (28)

1.03 F 0.37b 1.57 F 0.34 1.39 F 0.20

E51D(40) E477Q(40)

1.09 F 0.05 1.21 F 0.08

a

DOD

(kcat/Km)

DOD

(kcat/A)

0.32 F 0.28

DOD

(kcat/S0.5)

1.11 F 0.24

0.60 F 0.21b 1.35 F 0.62 1.01 F 0.24 0.57 F 0.13 1.37 F 0.20

Isotope effects (T = 25jC) are ratios (HOH/DOD) of pH- or pD-independent values obtained by least-squares fitting of rate parameters vs. pH or pD to simple models for bell-shaped profiles. b Entries for C221A were fit using a different procedure because there was insufficient data to get reasonable fits to bell-shaped profiles. Instead, isotopic ratios of parameters vs. pH or pD were fit to a model in which pKa estimates were fixed. The reported errors on these parameters are therefore minimum estimates of the uncertainty; more accurate error estimates would be larger. c The wild-type enzyme actually had a C-terminal His6 tag attached for efficient purification. The steady-state kinetics of the enzyme with and without the His tag are indistinguishable. d The C221 variants all have the C222A substitution as well.

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(C221) variants did not show sigmoidal kinetics, so simple Michaelis–Menten parameters (kcat and kcat/Km) were used. The values shown for the wild-type and C221 variants result from new fits of data published previously. Isotope effects reported recently (28) were not from least-squares fits but were estimates based on the apparent maxima in pH and pD rate profiles. It is now clear that the pH dependencies of the steady-state parameters are defined by narrow maxima that make analyses by inspection less reliable, although the earlier isotope effect estimates are within new and more accurate error limits. Results from rigorous least-squares fitting of the narrow pH/pD rate profiles produce error estimates that make detailed interpretations of our solvent isotope effects more difficult. Solvent isotope effects on kcat/B and kcat/ A are not reported for E51D and E477Q because the error estimates were very large; for the same reason, the isotope effect on kcat/B for wild-type YPDC is not shown in Table 1. The solvent isotope effects for the wild-type enzyme in Table 1 are in agreement with the values reported previously by Alvarez et al. (19) as 1.5 for kcat and 0.5 for kcat/A. B. Carbon Isotope Effects Carbon isotope effects were measured using a competitive method with natural-abundance levels of pyruvate-1-12C and pyruvate-1-13C. The product carbon dioxide was collected on a high-vacuum line at several fractional extents of the consumption of pyruvate and at 100% conversion of the substrate. The isotopic content of the carbon dioxide samples was determined by the isotopic mass spectrometry laboratory in the Geology Department at the University of Vermont. From these results, the kinetic isotope effect was calculated using the standard equation (30) for such competitive measurements. As shown in Table 2, these isotope effects report on either kcat/B for enzymes with sigmoidal steady-state kinetics or kcat/Km for C221A. The result for wild-type YPDC is consistent with several previous reports. O’Leary (31) reported a value of 1.0083 at pH 6.8; Jordan, Kuo, and Monse (32) found 1.0065 at pH 6.0; and the isotopic ratios reported by DeNiro and Epstein (33) correspond to 1.0063 at pH 6.0.

III. DISCUSSION A. Isotope Effects on kcat for Yeast Pyruvate Decarboxylase Variants All solvent isotope effects on kcat in Table 1 are smaller than expected for a single rate-limiting step with general acid or base catalysis involving generation of a strong hydrogen bond in the transition state. Alvarez et al. (18)


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Table 2 Carbon Kinetic Isotope Effects on Reactions Catalyzed by Yeast Pyruvate Decarboxylase and Variantsa 13

WT E477Q D28A C221Ac

(kcat/B)b

1.0046 1.0018 1.0398 1.0054

F F F F

0.0003 0.0009 0.0021 0.0004

a

Conditions were pH 6.0 at 25jC. Isotope effects are reported as the ratio of kcat/B for pyruvate-1-12C to pyruvate1-13C. Error limits are 95% confidence intervals for the standard deviation of the mean. c This variant also included a C222A substitution, and the isotope effect in this case is on kcat/Km. Source: Ref. 41. b

noted for the wild-type enzyme that the kcat solvent isotope effect was consistent with partial rate control by steps involving proton transfers after decarboxylation to facilitate acetaldehyde elimination. The similar size of the isotope effects for the three C221 variants and the fact that the turnover number is reduced by a factor of only about 2 suggests that there is little change in the rate-limiting steps and protonic aspects of the kcat reactant state(s) and transition state(s) on modifying the cysteine at the regulatory site. Collectively, the two active-site variants, E51D and E477Q, appear to have kcat solvent isotope effects that are slightly smaller than wild-type YPDC and the regulatory variants. The disruption of the active site may have altered the relative importance of several steps contributing to kcat to make proton transfer a smaller factor in the observed effect. Clearly, the active site variants do not show an increase in the solvent isotope effect, in spite of large decreases in turnover number, that would be expected if acid–base catalysis had become more rate limiting for kcat. The fact that the E51D isotope effect is significantly smaller than the values for E477Q and wild-type YPDC is consistent with a transition-state hydrogen bond between E51 and N1V of the thiamine diphosphate imino tautomer. The carbon isotope effects in Table 2 do not report directly on kcat. Instead, these competitive isotope effects only measure steps up to and in-

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cluding the first irreversible step, which is very likely the decarboxylation step. B. Isotope Effects at Low Pyruvate Concentrations for Pyruvate Decarboxylase Variants In the limit of infinitely low pyruvate concentration, the rate of wild-type YPDC will be reflected at the steady state in kcat/A and for the three regulatory C221 variants in kcat/Km. The inverse (faster in D2O than H2O) solvent isotope effect on kcat/A for wild-type YPDC has been attributed to the loss of the C221 SH proton on going from the free, unactivated enzyme to the rate-limiting transition state (19). The vibrational properties of the thiol group give it an isotopic fractionation factor that is close to 0.5 relative to water. If the thiol proton is present in the reactant state but absent in the transition state, Eq. (1) predicts a solvent isotope effect of 0.5. Taken together, the kcat/Km solvent isotope effects for the C221 mutants are consistent with this explanation. With the thiol removed from the 221 position, the solvent isotope effects are no longer inverse, although the uncertainties are large. In this explanation, the 0.5 reactant-state fractionation factor was assigned to the SH group of C221, but other assignments could be made consistent with the results. Infrared spectroscopic and isoelectric focusing studies of YPDC and its cysteine-substituted variants suggest the free enzyme at its optimal pH has C221 as a thiolate ion (34). With C221 as a thiolate rather than a thiol, the wild-type YPDC 0.5 fractionation factor could arise from a special hydrogen bond (35–37) located either near C221 or at a more remote position, perhaps at the active site, in communication with C221. For the active-site variants, E51D and E477Q, detailed interpretations of the solvent isotope effects for low-substrate conditions are not possible because the uncertainties in fitted pH- or pD-independent values for k/A and k/B were very large. We can instead make qualitative comparisons using k/S0.5, a parameter from fits to the Hill-type Eq. (2). The isotope effect of 0.57 for E51D as compared to 1.11 for wild-type PDC and 1.37 for E477Q can be interpreted in terms of an E51 transition-state hydrogen bond, perhaps to N1V of the thiamine diphosphate pyrimidine ring. Although it is difficult to assign specific reactant and transition states to k/S0.5, the trend in the isotope effects points to a transition-state fractionation factor in wild-type YPDC and E477Q that more than compensates for the 0.5 reactant-state fractionation factor associated with the regulatory site. The carbon isotope effects for wild-type YPDC and the active-site variants correspond to YPDC at substrate concentrations lower than saturating conditions but not the infinitely low substrate condition of kcat/A. Instead, the carbon effects report on kcat/B, the kinetic parameter defined by a reactant state after the first molecule of pyruvate binds to the enzyme, leaving


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the remaining equivalent of substrate free in solution. For the regulatory-site variants with simple Michaelis–Menten kinetics, the carbon isotope effects correspond to kcat/Km. The reactant-state fractionation factor [Eq. (1)] for all of the carbon isotope effects is therefore the same for all of our carbon isotope effects and measures the preference for 13C over 12C at the free pyruvate carboxyl carbon. The shifts in the carbon isotope effects on going from wild-type YPDC to either of the regulatory variants can be explained in terms of changes in rate-limiting steps. If two steps in series contribute to the observed rate constant, the isotope effect will be a weighted average of the isotope effects on the net rate constants corresponding to the two steps. The reactant-state fractionation factor [Eq. (1)] for both steps will be the same free pyruvate value, while the transition-state fractionation factors will depend on the structure around the carboxylate carbon at the barrier heights. For steps that precede the decarboxylation step, the carboxylate closely resembles the structure in free pyruvate, so the ratio of transition-state and reactant-state fractionation factors will be very close to unity. For the decarboxylation step, the ratio of fractionation factors should be close to 1.05, because the C–C stretching mode is lost in the transition state. The observed carbon isotope effect should therefore appear as a weighted average of these two limits [Eq. (4)], with the weighting factors w and (1 w) determined by the free-energy difference between the two transition states [Eq. (5)]: ðk=BÞ12 ¼ 13 ðk=BÞ ¼ wð1:05Þ þ ð1 wÞð1:00Þ ðk=BÞ13 TS w Gdecarboxylation GTS other step ln ¼ 1w RT

ð4Þ ð5Þ

As was noted in Section I, decarboxylation is not the main rate-limiting step for k/B of wild-type YPDC. Using our observed carbon isotope effect of 1.0046 and Eq. (4), decarboxylation contributes only 9% to k/B, and the decarboxylation transition-state free energy is 5.8 kJ/mol less than that of a higher-energy transition state that precedes decarboxylation. Our results show that the active-site variants shift the relative importance of the two transition states. E477Q has a slightly smaller carbon isotope effect of 1.0018, indicating even less rate control by decarboxylation than was seen for wildtype PDC, while D28A has a much larger isotope effect of 1.0398, consistent with decarboxylation as the dominant rate-limiting step for this variant. We conclude, therefore, that D28 plays a more important role in stabilizing the transition state for decarboxylation than does E477. In contrast to the active-site variants, the regulatory variants appear not to shift the relative transition-state energies from the wild-type positions because the carbon isotope effects on k/B for C221A/C222A, C221E/C222A,

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and C221D/C222A are unchanged from the wild-type value. The disruptions introduced at the remote regulatory site translate into larger overall energies of activation, but the relative destabilization of the rate-limiting transition states up to and including decarboxylation must be by the same energetic amount. This suggests that the ultimate effect of C221 in conjunction with pyruvate bound in the regulatory site is to stabilize transitition states uniformly, perhaps through adjustments in the protein to relieve common unfavorable steric interactions. C. Conclusions The active-site variants of YPDC decrease the low-substrate activity of the enzyme by selectively destabilizing transition states in the decarboxylation phase of the catalytic cycle. Because the carbon isotope effect increases dramatically when D28 is modified, this residue is especially important in stabilizing the decarboxylation transition state. Other studies have shown that D28 and E477 are also an important contributors to transition-state stabilization in steps after decarboxylation (38). Solvent kinetic isotope effects under low-substrate conditions are consistent with a reactant-state site having / f0.5 located near C221. Under kcat conditions, the solvent isotope effects provide support for partial rate control by general acid- or base-catalyzed processes, featuring low-ø hydrogen bonds to E51 in transition states.

ACKNOWLEDGMENTS This research was supported by NIH grant GM50380. REFERENCES 1. 2. 3.

4. 5.

R Kluger. Thiamin diphosphate: a mechanism update on enzymic and nonenzymic catalysis of decarboxylation. Chem Rev 87:863–876, 1987. RL Schowen. Thiamin-dependent enzymes. In: M Sinnot, ed. Comprehensive Biological Catalysis, vol. 2. London: Academic Press, 1998, pp 217–266. F Jordan. Interplay of organic and biological chemistry in understanding coenzyme mechanisms: example of thiamin diphosphate–dependent decarboxylations of 2-oxo acids. FEBS Lett 457:298–301, 1999. F Jordan. Current mechanistic understanding of thiamin diphosphate–dependent enzymatic reactions. Natural Product Reports 2003 issue 2, in press. G Hu¨bner, S Ko¨nig, A Schellenberger. The functional role of thiol groups of pyruvate decarboxylase from brewer’s yeast. Biomed Biochim Acta 47:9–18, 1988.

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I Barburina, Y Gao, Z Hu, F Jordan. Substrate activation of brewer’s yeast pyruvate decarboxylase is abolished by mutation of cysteine 221 to serine. Biochemistry 33:5630–5635, 1994. F Jordan, N Nemeria, F Guo, I Baburina, Y Gao, A Kahyaoglu, H Li, J Wang, J Li, JR Guest, W Furey. Regulation of thiamin diphosphate–dependent 2-oxo acid decarboxylases by substrate and thiamin diphosphate Mg(II)—evidence for tertiary and quaternary interactions. Biochim Biophys Acta 1385:287–306, 1998. P Arjunan, T Umland, F Dyda, S Swaminanthan, W Furey, M Sax, B Farrenkopf, Y Gao, D Zhang, F Jordan. Crystal structure of thiamin diphosphate–dependent enzyme pyruvate decarboxylase from the yeast Saccharomyces cerivisiae at 2.3-A˚ resolution. J Mol Biol 256:590–600, 1996. Chapter 5 in the present volume. MM Kreevoy, DG Truhlar. Transition-state theory. In: CF Bernasconi, ed. Investigation of Rates and Mechansims of Reactions, 4th ed. New York: Wiley, 1986. Part 1, pp 13–95 S Hammes-Schiffer. Comparison of hydride, hydrogen atom, and proton-coupled electron transfer reactions. Chem Phys Chem 3:33–42, 2002. KB Schowen, RL Schowen. Solvent isotope effects on enzyme systems. Meth Enzymol 87:551–605, 1982. DM Quinn, LD Sutton. Theoretical basis and mechanistic utility of solvent isotope effects. In: PF Cook, ed. Enzyme Mechanism from Isotope Effects. Boca Raton, FL: CRC Press, 1991, ch 3. EH Cordes. Mechanism and catalysis for the hydrolysis of acetals, ketals, and ortho esters. Prog Phys Org Chem 4:1–44, 1967. SS Minor, RL Schowen. One-proton solvation bridge in intramolecular carboxylate catalysis of ester hydrolysis. J Am Chem Soc 95:2279–2281, 1973. R Eliason, MM Kreevoy. Kinetic hydrogen isotope effects in the concerted mechanism for the hydrolysis of acetals, ketals, and ortho esters. J Am Chem Soc 100:7037–7041, 1978. WP Huskey. Origins and interpretations of heavy-atom isotope effects. In: PF Cook, ed. Enzyme Mechanism from Isotope Effects. Boca Raton, FL: CRC Press, 1991, ch 2. FJ Alvarez, J Ermer, G Hu¨bner, A Schellenberger, RL Schowen. Catalytic power of pyruvate decarboxylase. Rate-limiting events and microscopic rate constants from primary carbon and secondary hydrogen isotope effects. J Am Chem Soc 113:8409–9402, 1991. FJ Alvarez, J Ermer, G Hu¨bner, A Schellenberger, RL Schowen. The linkage of catalysis and regulation in enzyme action. Solvent isotope effects as probes of protonic sites in the yeast pyruvate decarboxylase mechanism. J Am Chem Soc 117:1678–1683, 1995. F Jordan, YH Mariam. N1V-Methylthiaminium diiodide. Model study on the effect of a coenzyme-bound positive charge on reaction mechanisms requiring thiamin pyrophosphate. J Am Chem Soc 100:2534–2541, 1978. A Schellenberger. The amino group and steric factors in thiamin catalysis. Ann N Y Acad Sci 378:51–62, 1982.

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22. F Jordan, G Chen, S Nishikawa, B Sundoro-Wu. Potential roles of the aminopyrimidine ring in thiamin-catalyzed reactions. Ann N Y Acad Sci 378: 14–31, 1982. 23. F Jordan. Role of the aminopyridine ring in thiamin-catalyzed reactions. II. Proton NMR evidence for high barriers to amino group rotation in 4-aminopyrimidines, including thiamin, at low pH in water. J Org Chem 47:2748–2753, 1982. 24. F Jordan, Z Zhang, E Sergienko. Spectroscopic evidence for participation of the 1V,4V-imino tautomer of thiamin diphosphate in catalysis by yeast pyruvate decarboxylase. Bioorg Chem 30:188–198, 2002. 25. M Killenberg-Jabs, S Ko¨nig, I Eberhardt, S Hohmann, G Hu¨bner. Role of Glu51 for cofactor binding and catalytic activity in pyruvate decarboxylase from yeast studied by site-directed mutagenesis. Biochemistry 36:1900–1905, 1997. 26. EA Sergienko, F Jordan. Catalytic acid–base groups in yeast pyruvate decarboxylase. 3. A steady-state kinetic model consistent with the behavior of both wild-type and variant enzymes at all relevant pH values. Biochemistry 40:7382– 7403, 2001. 27. I Barburina, Y Gao, Z Hu, F Jordan. Substrate activation of brewer’s yeast pyruvate decarboxylase is abolished by mutation of cysteine 221 to serine. Biochemistry 33:5630–5635, 1994. 28. W Wei, M Liu, F Jordan. Solvent kinetic isotope effects monitor changes in hydrogen bonding at the active center of yeast pyruvate decarboxylase concomitant with substrate activation: the substituent at position 221 can control the state of activation. Biochemistry 41:451–461, 2002. 29. R Hill. Chemical nature of hemochromogen and its carbon monoxide compound. Proc Roy Soc (London) 100B:419–430, 1926. 30. MH O’Leary. Determination of heavy-atom isotope effects on enzyme-catalyzed reactions. Meth Enzymol 64B:83–104, 1980. 31. MH O’Leary. Carbon isotope effect on the enzymatic decarboxylation of pyruvic acid. Biochem Biophys Res Commun 73:614–618, 1976. 32. F Jordan, DJ Kuo, EU Monse. Carbon kinetic isotope effects on pyruvate decarboxylation catalyzed by yeast pyruvate decarboxylase and models. J Am Chem Soc 100:2872–2878, 1978. 33. MJ DeNiro, S Epstein. Mechanism of carbon isotope fractionation associated with lipid synthesis. Science 197:261–263, 1977. 34. I Barburina, DJ Moore, A Volkov, A Kahyaoglu, F Jordan, R Mendelsohn. Three of four cysteines, including that responsible for substrate activation, are ionized at pH 6.0 in yeast pyruvate decarboxylase: evidence from Fourier transform infrared and isoelectric focusing studies. Biochemistry 35:10249– 10255, 1996. 35. MM Kreevoy, TM Liang. Structures and isotopic fractionation factors of complexes, A1HA21. J Am Chem Soc 102:3315–3322, 1980. 36. WW Cleland. Low-barrier hydrogen bonds and low-fractionation-factor bases in enzymatic reactions. Biochemistry 31:317–319, 1992.


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37. WP Huskey. Model calculations of isotope effects using structures containing low-barrier hydrogen bonds. J Am Chem Soc 118:1663–1668, 1996. 38. EA Sergienko, F Jordan. Catalytic acid–base groups in yeast pyruvate decarboxylase. 2. Insights into the specific roles of D28 and E477 from the rates and stereospecificity of formation of carboligase side products. Biochemistry 40: 7369–7381, 2001. 39. J Wang, R Golbik, B Seliger, M Spinka, K Tittmann, G Hu¨bner, F Jordan. Consequences of a modified putative substrate-activation site on catalysis by yeast pyruvate decarboxylase. Biochemistry 40:1755–1763, 2001. 40. M Liu. Studies of Yeast Pyruvate Decarboxylase. 1. Function of the Active Center Acid-Base Groups. 2. Partitioning of Thiamin-Bound Covalent Intermediates. PhD dissertation, Rutgers University—Newark, 2002. 41. L Chen, Y Yuan, WP Huskey. Submitted for publication. 42. PJ Kraulis. MOLSCRIPT: A program to produce both detailed and schematic plots of protein structures. J Appl Cryst 24:946–950, 1991.

14 Insights into the Mechanism and Regulation of Bacterial Acetohydroxyacid Synthases David M. Chipman, Ze’ev Barak, Stanislav Engel, Sharon Mendel, and Maria Vyazmensky Ben-Gurion University of the Negev, Beer-Sheva, Israel

I. INTRODUCTION Acetohydroxyacid synthase (AHAS) is a member of a homologous family of TDP-dependent enzymes whose initial step is decarboxylation of pyruvate or another 2-ketoacid (1,2), such as pyruvate oxidase and benzoyl formate decarboxylase. Many representatives of this family are discussed in this book. However, despite the similarity in sequence and cofactor requirements of bacterial AHASs to, e.g., pyruvate oxidase from Lactobacillus plantarum (LpPOX) (3–5), they have unique properties of considerable interest. The biosynthetic AHASs catalyze decarboxylation of pyruvate, followed by the specific condensation of the bound hydroxyethylTDP (HETDP) intermediate with a second aliphatic ketoacid to form an acetohydroxyacid; this reaction is the first common step in the pathway for the biosynthesis of the branched-chain amino acids (6) (Fig. 1). The partition of the flux through AHAS between acetohydroxybutyrate and acetolactate determines the relative rates of formation of isoleucine and of valine, leucine, and the coenzyme A precursor pantothenate, respectively (7) (Fig. 1). In order to allow the pathway to function with an intracellular concentration of 2233

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Figure 1 The reaction catalyzed by AHAS. The decarboxylation of pyruvate to form the enzyme-bound anion/eneamine of hydroxyethylthiamine diphosphate (HETDP) is followed by condensation with a second keto acid and release of an acetohydroxyacid. AHB is converted to isoleucine by the action of three additional enzymes; the same three enzymes can convert acetolactate to valine. The immediate precursor of valine is also a precursor for biosynthesis of leucine and pantothenate.

ketobutyrate (the precursor of acetohydroxybutyrate), almost two orders of magnitude lower than that of pyruvate (8), most AHASs show a very strong preference for 2-ketobutyrate as the second substrate (9). The biosynthetic AHASs require bound FAD as a cofactor, despite the fact that the physiological AHAS reaction does not involve any redox step (10). Structurally, bacterial AHASs are hetero-oligomers; in addition to the catalytic subunits homologous to the related TDP-dependent enzymes, AHASs have unique regulatory subunits. Finally, most bacterial AHASs show feedback inhibition by one or more branched-chain amino acids; the most common pattern is mixed V- and K-type inhibition by valine, as seen, for instance, in Escherichia coli isozyme III. We describe here some recent findings in our laboratories on AHAS II and III from E. coli, focusing on two aspects of bacterial AHASs that are crucial to understanding their physiological functions: substrate specificity and regulation. II. CATALYSIS AND SPECIFICITY A. The First Step The role of the AHAS II active site in promoting the formation of HETDP from pyruvate is now quite clear. Figure 2 shows some of the residues in the

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Figure 2 Hypothetical model for the active site of AHAS II, in the process of condensation of 2-ketobutyrate with HETDP. TDP is shown with dark, thick bonds in the lower center of the picture, and the substrate-derived atoms with thinner, white bonds. Side chains of residues of interest are shown; residues from one subunit have primed numbers and darker bonds than those from the other. The grey ellipse outlines a region whose structure is less well defined at this time. The arrow indicates a probable path of substrate molecules from the solvent to and from the active site.

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region around TDP in our model of E. coli AHAS II, which was based on the crystal structure of the homologous enzyme LpPOX (5). The recently published crystal structure of a dimer of catalytic subunits of yeast AHAS (11) confirms that the thiamine cofactor occupies a site very similar to that in LpPOX. The required divalent cation is bound between the pyrophosphate moiety of TDP and the side chains of conserved Asp and Asn residues in a ‘‘TDP motif,’’ the carboxyl group of a conserved glutamate is within hydrogen-bonding distance of N-1 of the aminopyrimidine, and a conserved methionine is wedged between the thiazole and pyrimidine rings, enforcing the V-conformation. Site-directed mutagenesis of AHAS II has verified the functional role of the Asp428 side chain in binding Mg2+ and of Glu47 as a critical part of the relay system involved in deprotonation at C-2 of the thiazolium (12). Mutagenesis of Met403 (Vitaly Balan, unpublished results) shows that this residue also makes an important contribution to TDP binding. It is reasonable to assume that the first steps in AHAS catalysis, i.e., decarboxylation of pyruvate and formation of HETDP, proceed much as they do in related enzymes. It should be pointed out, however, that in AHAS II the formation of HETDP is quite slow (f24 s1) as compared to other enzymes for which this reaction has been measured (Chapter 5, this volume). B. The Product-Determining Second Step 1. Specificity Unfortunately, it is much more difficult to suggest roles for protein residues in the final steps of the reaction in which the product is determined, because the structure of the region we believe is critical for selective recognition of the second ketoacid is far less clear. Part of the region, circled in Figure 2, is very different in sequence from the equivalent region in LpPOX, so modeling with great confidence is not possible. The equivalent region in the published crystal structure of the yeast enzyme is disordered (11). We showed several years ago (without purification of the proteins in question) that mutagenesis of a single amino acid in this region, Trp464, leads to a drastic reduction in the preference of AHAS II for 2-ketobutyrate as the second substrate (5). We refer to this preference, characteristic of a given AHAS, as ‘‘R,’’ where ðAHB formedÞ=ðAL formedÞ ¼ R ½2 ketobutyrate=½pyruvate On the strength of this observation, we suggested that the indole ring of Trp464 makes contact with the C-4 methyl group of 2-ketobutyrate at the transition state(s) for the second stage of the reaction. The orientation of the indole ring and of the bound product shown in Figure 2 were chosen accordingly.


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We have since repeated and expanded these mutagenesis experiments, having expressed the wild-type and mutant proteins at high levels and purified them to near-homogeneity. Several observations are significant (Table 1): Replacement of Trp464 with any other amino acid (including Phe and Tyr, not shown) leads to at least an order-of-magnitude decrease in R, but in many cases the catalytic activity of the mutant enzyme is close to that of the wild type. No other residue in this region seems to have this kind of effect on substrate specificity. Trp464 also has a striking effect on inhibition by the sulfonylurea herbicide SMM (Table 1), but it is only one of many sites that have strong effects on the herbicide sensitivity of the enzyme (reviewed in Refs. 13 and 14). For comparison, Table 1 shows the effects of some mutations at Val375 and Val461 (15). These residues presumably lie on either side of the thiazole ring, not far from C-2. Given this, replacing them by Ile would introduce some crowding. V-I mutations at either—or both—positions have, however, relatively small effects on the enzyme activity. The mutation V375I, quite close to the thiazolium, reduces the apparent affinity for TDP by 4.5-fold, with few other effects. The mutation V461I, in the general region where we think the second substrate is bound, decreases kcat/Km by 30-fold but only reduces the specificity for 2-ketobutyrate as second substrate by less than threefold. This mutation also has a small but significant effect on SMM sensitivity. The double mutant V375I–V461I shows the combined effects of the two separate mutations (Table 1).

Table 1 Properties of AHAS II and Some Mutants at Trp464, Val375, and Val461a Enzyme Wild-type W464L W464E V375I V461I V-I doubleb Specific activity, U/mg kcat/KM(pyr), M1s1 Rc K0.5, TDP, lM Ki, SMM, lM d a

20 9000 58 0.6 0.9

17 2400 2.2 2.2 350

1.5 190 2.2 0.7 7000

16 7600 60 2.8 0.8

2.2 280 22 0.5 3.6

2.85 390 23 2.7 3.4

Activity measurements with purified AHAS II, in a standard colorimetric assay in the presence of 50 mM pyruvate, in 0.1 M potassium phosphate buffer at pH 7.6 (Ref. 12). b Double mutant V375I–V461I. c R is the specificity ratio, where (AHB formed)/(AL formed) = R [2-ketobutyrate]/[pyruvate] d Ki for SMM is the concentration for apparent half-inhibition in the foregoing single-point colorimetric assay.

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2. Methionine 250 Mutation of Met250 leads to an interesting and completely unexpected effect (Fig. 3). With wild-type AHAS II and many other mutants of this enzyme (as well as at least four other AHASs), the competition between pyruvate and 2-ketobutyrate does not effect the total rate of formation of acetohydroxy acids (Fig. 3A). As the concentration of 2-ketobutyrate is increased in the presence of a fixed pyruvate concentration, acetohydroxybutyrate replaces acetolactate as product, but the sum of the rates of formation of the two remains essentially constant (9,16). Such behavior requires that the rate-determining step for enzyme turnover be different from the product determining step (16). Unless the rate constants of parallel steps with pyruvate and 2-ketobutyrate as second substrate are coincidentally identical, it further implies that the rate-determining step occurs prior to binding of the second substrate. With mutant M250A, in contrast, the total rate of product formation increases by more than sevenfold upon addition of 2-ketobutyrate (Fig. 3B). A similar, but less striking, effect is seen with M250L (not shown). The

Figure 3 Competition between acetolactate formation and acetohydroxybutyrate formation, for wild-type AHAS II (A) and the Met250Ala mutant (B). In each case, the reaction was carried out under standard conditions in the presence of 50 mM pyruvate, with a varied amount of 2-ketobutyrate. After six minutes, the formation of acetolactate (.) and acetohydroxybutyrate (E) was measured simultaneously by GLC analysis. The sum of the two products is also shown (o).


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apparent specificity, R, of each of these mutants is close to that of the wildtype. The values of kcat/Km with pyruvate as sole substrate are 130 and 875, respectively, for M250A and M250L. For these mutants then, the second substrate is involved in the rate-determining step for turnover; the mutation has presumably changed the rate-determining step by lowering the rate constant for one or more steps in the turnover cycle. Which step in the process could conceivably be affected by modification of this nonpolar residue? The recent results of Hu¨bner’s enzymology group at Martin Luther University in Halle, Germany, allow us to speculate in this regard. They applied their NMR technique for assessing the distribution of TDP-containing intermediates in an enzymatic reaction at steady state to a hexahistidine-fused wild-type AHAS II and obtained the estimates for individual rate constants, summarized in Figure 4 (this volume, Chapter 5; K. Tittmann, personal communication, 2002). The slowest step(s) in the reaction by far is the formation of the first covalent adduct, lactyl-TDP. It is thus not surprising that the turnover rate for wild-type AHAS II is unaffected by the competition between second substrates. It may be significant that in the presence of 2-ketobutyrate, the last two steps (i.e., formation of the acetohydoxybutyrate–TDP adduct and release of the product) are estimated to have rate constants greater than 2000 s1. The kinetic effects of replacement of Met250 could be explained if the mutation lowered the rate of either or both of the last two steps to a rate equal to or lower than that of lactyl–TDP formation. Unless Met250 is much closer to the bound HETDP than suggested by our model (Fig. 2), it seems unlikely that this residue has a role in the covalent bond–making reaction between the intermediate and the second substrate. Met250 is more likely to be in the substrate access channel; the homologous Met354 residue in the yeast enzyme is in such a region (11). (In this regard, it should be noted that several nonconservative mutations at Met354 in the yeast enzyme lead to an active, herbicide-resistant enzyme (17).) We suggest, therefore, that Met250 has an effect on access to and from the active site and thus on the product-release step. We hope that direct measurements of TDP intermediates on M250A and other mutants will allow us to test this suggestion. 3. Polar Side Chains We have mutated each of the residues of AHAS II shown in Figure 2, usually making two or three different mutations at a given position. Two of these residues, Gln110 and His251, have polar side chains and are in regions where they could conceivably interact with oxygen atoms of the nascent TDP-bound product. Mutant enzymes modified at Gln110 (Q110A, Q110N, and Q110H) have values of kcat/Km in the reaction with pyruvate that are 30- to 100-fold

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Figure 4 Minimal mechanism for AHAS. The rate of deprotonation of bound TDP was determined by H/D exchange in the absence of substrate (12). The other pseudo-first-order rate constants shown were estimated for reaction of wild-type AHAS II in the presence of a saturating concentration of pyruvate as sole substrate, from kcat for the enzyme and the partition of TDP among the several intermediates (this volume, Chapter 5; K. Tittmann, personal communication, 2002).

lower than that of the wild-type enzyme. The side chain of Gln110 is probably close to the 4V-amine nitrogen of TDP (the homologous residues Gln202 in yeast AHAS (11) and Gln 122 in LpPOX (4) are less than 3.4 A˚ from this atom in the crystal structures) and may thus affect multiple steps of the reaction. Mutations at His251 have a more modest effect; H251Q and H251A have values of kcat/Km that are 8- and 11-fold lower, respectively, than that of the wild-type enzyme. To clarify the mechanistic roles of these polar residues, it will be necessary to examine the effects of mutations on the distribution of intermediates or on partial reactions of the enzyme.


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Figure 5 The reaction of pyruvate and benzaldehyde to form R-phenyl acetyl carbinol.

C. A New Condensation Reaction Catalyzed by AHAS AHASs are quite specific in catalyzing a condensation of a second ketoacid with the bound HETDP intermediate to form S-acetohydroxyacids (Fig. 1). Alternative reactions, such as protonation and release of acetaldehyde or electron transfer to the bound FAD, are the major fates of the intermediate in some related enzymes, yet they occur very slowly in AHAS. However, we have

Figure 6 Stereochemistry of the formation of S-acetolactate and R-PAC by AHAS II.

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found that AHAS II can catalyze a novel nonphysiological reaction, condensation of HETDP with benzaldehyde to form R-phenyl acetyl carbinol (PAC) (Fig. 5) with good efficiency. This reaction, which can be extended to other AHASs and other aromatic aldehydes, is of considerable practical interest for chiral syntheses (patent pending). Here we will focus on an interesting mechanistic implication of this reaction. When a mixture of pyruvate and benzaldehyde reacts in the presence of AHAS II, the relative initial rate of formation of acetolactate and PAC depends, as expected, on the ratio between the two reagents. At a given concentration of pyruvate, the formation of PAC increases with increasing benzaldehyde levels, at the expense of acetolactate formation, as expected if the two ‘‘second substrates’’ compete for a common intermediate. If we make the reasonable assumption that the carbonyl C j O bond of the second substrate has the same orientation in the active site in the two alternative reactions, then the stereochemistry of the products requires that the phenyl group of benzaldehyde and the carboxylate of pyruvate occupy similar regions in the enzyme active site (Fig. 6). The stereospecificity of the two reactions would thus suggest that it may be an oversimplification to search for a protein polar side chain as a central element in recognition of the carboxylate of a second ketoacid substrate. III. FEEDBACK INHIBITION BY BRANCHED-CHAIN AMINO ACIDS Valine inhibition of AHAS plays a major role in the physiological control of the biosynthesis of the three branched-chain amino acids in bacteria. Escherichia coli and other enterobacteria encode three different isozymes of AHAS that differ in their response to valine. AHAS II, discussed earlier, is unusual in that it is insensitive to valine inhibition. The valine-sensitive AHAS III, on the other hand, is more typical of bacterial AHASs. The valine-binding site of AHAS III is located in the small (17-kDa) regulatory subunit (SSU) required for full catalytic activity and valine sensitivity of the enzyme (18–20). The isolated large (62-kDa) catalytic subunit has 3–5% of the activity of the holoenzyme and is not sensitive to feedback inhibition, while the isolated SSU binds valine and has no catalytic activity of its own (21). AHAS III can be completely reconstituted from its subunits (21). The valine inhibition of the enzyme differs from the classical allosteric model. The substrate dependence of AHAS III is noncooperative in both the presence of and the absence of valine, while the observed effect of valine is of a mixed V/K type. The enzyme has an apparent Km for pyruvate about 3.2-fold higher when saturated with valine than in its absence, and an apparent kcat about 1.6-fold lower. The final inhibition level is dependent on pyruvate, so at


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100 mM pyruvate, the enzyme is about 50% inhibited at saturating valine concentrations, while at a typical intracellular pyruvate concentration of 0.5 mM, the enzyme is 80% inhibited at saturation (22). It has been proposed in several cases that an allosteric inhibitor of an enzyme might affect kcat by restricting the dynamics of protein domain movements required for turnover, such as the opening and closing of a substrate binding cleft (23–25). We already noted that for AHAS II, the catalytic rate is largely determined by the formation of the first covalent intermediate, lactyl-TDP. This step is, in fact, far slower than the rate of the same process in pyruvate decarboxylase or pyruvate oxidase (this volume, Chapter 5; K. Tittmann, personal communication, 2002). This process might require closing of the active site cleft, in a relatively slow process that could be restricted by binding of the allosteric inhibitor to the SSU in AHAS III. A. Structure of the Valine Site We made a first step toward a structural explanation of the regulation of AHAS by analyzing some spontaneous valine-resistant mutants of AHAS III (21,22). The location of several of these mutations in the regulatory subunit of AHAS III was in very good agreement with a model based on a fold-recognition algorithm (22). The model suggests that a 76 amino acid N-terminal domain of the SSU may fold like the C-terminal serine-binding regulatory domain (24) of 3-phosphoglycerate dehydrogenase (3PGDH). This fold is an a-h sandwich with a ferredoxin-like hahhah topology (Fig. 7), which has recently been recognized as characteristic of a family of protein domains that includes many regulatory ligand-binding domains (26,27). Pairs of regulatory domains interact in the quaternary structure of 3PGDH to form two symmetry-related ligand-binding sites in the interface between them (24). The properties of a number of Val-resistant mutants of AHAS III provide strong support for the idea that the N-terminal domain of the AHAS III SSU folds and forms pairs of valine sites in a homologous manner (22). Even if the valine-binding site of AHAS III indeed has the same topology as the serine-binding site of 3PGDH (Fig. 7), the detailed interactions that lead to ligand recognition must of course be different. Serine makes polar interactions with two asparagine side chains in 3PGDH that are conserved in the AHAS III SSU (N11 and N29), but both serine carboxyl oxygens interact with a histidine residue that is not conserved (23) (Fig. 8A). The residue in the homologous position in the AHAS III SSU is Leu9, which is surrounded by a hydrophobic region (Fig. 8B). The carboxyl group of valine must thus point toward some polar group in a different part of the site,

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Figure 7 Ribbon diagram of a symmetric dimer of 3PGDH C-terminal regulatory domains, which form a pair of serine-binding sites. The diagram was prepared from the published coordinates of Ref. 30, using the program Molscript (Ref. 31). The large spheres represent the regulatory ligand serine. We suggest that the N-terminal domain of the AHAS III SSU folds in a similar way, with valine in the ligand-binding sites (Ref. 22).

perhaps one of the conserved asparagine residues Asn11 or Asn29. A series of mutations of residues around the putative valine-binding site (Table 2) support the binding mode for valine suggested in Fig. 8B. We suggest that the amino group of valine, rather than its carboxyl, is close to Asn11, because replacement of the carboxamido group of Asn11 with a carboxylate (N11D) leads to an enzyme that is clearly valine sensitive, albeit with a Ki 10-fold higher. Some valine-sensitive AHAS SSUs (e.g., from Corynebacterium glutamicum) have an aspartate residue at this position. Replacement by aspartate of Asn29, which is completely conserved in valine-sensitive SSUs, causes a much greater loss of valine sensitivity.


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Figure 8 (A) Closer view of one of the ligand sites of 3PGDH (see Fig. 7), showing the polar contacts of bound serine. These are between the amine nitrogen of serine and both Asn346 of one chain and Asn364 of the other, between the two carboxyl oxygens and His344, and between the side-chain hydroxyl and bound waters. (B) Hypothetical binding of valine in a ligand site of AHAS III. (See text.)

B. The C-Terminal Domain of the Regulatory Subunit The proposed N-terminal domain described earlier includes only about 76 out of the 163 amino acid residues of the SSU polypeptide. To test the hypothesis that this stretch of amino acids acts as a discrete domain of the SSU, as well as to examine the role of the remaining C-terminal half of the polypeptide, we prepared a series of truncations from the carboxyl end of the SSU: D35, D48, D80, D95, and D112. The D35, D48, D80, and D95 constructs all lead to essentially complete activation of the catalytic subunit, although their apparent affinities for the large subunit vary (Table 3). The D80 construct, corresponding to a few more amino acid residues than the putative N-

Table 2 Valine Sensitivity of AHAS III Reconstituted Using Wild-Type and Mutant SSUs SSU Ki, lM

Wild-type 11

N11A >5000

N11D 200

N29H 2200

N29D 1000

L9V 22

L9H 3600

Q59L 32

G14A 3900

Purified AHAS III large subunit was reconstituted with a 10-fold excess of the SSU in question and the activity at varying valine concentrations measured in the presence of 50 mM pyruvate in KPi buffer at pH 7.6. The apparent valine inhibition constant Ki was determined by fit of the initial velocity data to V = Vf + Ki (V0 Vf)/([Valine] + Ki).

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Table 3 Properties of Wild-Type and Truncated AHAS III Small Subunits Small subunit Wild type D35 D48 D80 D95

Relative reconstituted activitya 1 1.09 1.1 1.35 f1

SSU concentration for 50% activation, lM a

Valine inhibition Ki, mM b

Valine-binding Kd, mM c

0.27 0.64 0.6 0.42 24

0.037 F 0.008 z2 z2 z2 z2

0.15 >20 >20 >20 >20

AHAS III large subunit (0.088 lM ) was preincubated with varying quantities of wild-type or truncated SSU and the AHAS activity assayed by the standard procedure (Ref. 22). The activation was approximated by the Hill equation, and this fit was used to estimate the maximal activity (expressed relative to that obtained with wild-type SSU) and the concentration of SSU for half-activation. b AHAS activity was measured with a 15- to 20-fold molar excess of SSU in the presence of 10 mM pyruvate, under standard conditions (Ref. 22) with different valine concentrations. The observed activity was fit to an equation for partial inhibition: V = Vf + (V0 Vf)Ki/([Valine] + Ki). For the wild-type, Vf was f37% of V0 (i.e., 63% inhibition). Vf could not be estimated well for any of the truncated mutants. c The binding of valine to the SSU alone was measured by equilibrium dialysis in a buffer containing 0.9 M tricine and 0.25 M MgCl2, at pH 8.5. a

terminal domain, has the highest affinity for the catalytic subunit and leads to a reconstituted enzyme with kcat/KM about 35% higher than that of the wildtype enzyme. The D35 and D48 SSUs have somewhat lower affinities for the large subunit. The D95 SSU, which corresponds to the putative N-terminal domain with its last h-strand removed, has an affinity for the catalytic subunit nearly two orders of magnitude lower than that of the wild-type or the truncated constructs, which include the whole of the domain (Table 3). The D112 construct did not accumulate at all in the host cells used to produce it; we assume that it was rapidly degraded. These results provide further clear support for the proposal that the AHAS III SSU has a discrete N-terminal domain some 75–80 amino acid residues long that can fold independently. This domain is sufficient for recognition and activation of the catalytic subunit. On the other hand, none of the truncated constructs shows the valine inhibition characteristic of the enzyme reconstituted with the full-length SSU (Table 3). In addition, while the isolated wild-type SSU can be shown to bind valine (21,28) (albeit with an affinity considerably lower than expected from Ki), none of the truncated constructs binds valine. We have recently succeeded in measuring equilibrium binding of valine to the reconstituted wild-


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type holoenzyme (Kd f 0.01 mM), but similar experiments with the active enzyme reconstituted with the D80 construct suggest that it binds valine 50 times more weakly, if at all. Thus, while the N-terminal portion of the SSU is sufficient for activation of the catalytic subunit, the C-terminal portion of the SSU is required for the formation of valine-binding sites and for the regulatory response. IV. CONCLUSIONS AND SPECULATIONS The catalytic specificity of AHAS is due to a number of factors, many of which we understand only poorly. The intermediate HETDP is not protonated rapidly, presumably because the active site excludes solvent and has no effective proton-donating groups near the eneamine/anion. Duggleby and coworkers suggest that in AHAS, electrons are not transferred rapidly from HETDP to FAD because of the planar conformation of the flavin system (11). What is more important is that HETDP is formed slowly but reacts rapidly with a second substrate (Fig. 4), so at steady-state less than 3% of the enzyme is in the form of HETDP (This volume, Chapter 5; K. Tittmann, personal communication, 2002). The important question, then, is why the condensation of the HETDP with a second ketoacid is so fast (Fig. 4): If the reaction with, e.g., 2-ketobutyrate, is in fact a second-order process, it must have a rate constant greater than 4 104 M1s1. It is, therefore, tempting to speculate that the second substrate can bind before the decarboxylation step so that the condensation would be a firstorder process. On the other hand, in AHAS II the binding of the second 2ketoacid cannot be absolutely required for decarboxylation, since pyruvate or 2-ketobutyrate competes with oxygen for HETDP (29). As we have seen, the nature of the bound second substrate does not affect the rate determining step in the wild-type AHAS. It could, however, affect more than one of the subsequent steps, so the origin of second substrate specificity remains a complex question. Examination of the distribution of intermediates in some mutated AHASs with altered specificity (e.g., W464 mutants in AHAS II) or altered rate-determining steps (M250 mutants) may help shed further light on this question. The possibility that the slow process leading to formation of the first covalent intermediate in the AHAS reaction (Fig. 4) involves a significant conformational change in the enzyme provides a possible mechanism for Vtype allosteric inhibition. The work we describe identifies a valine-binding domain of the AHAS III regulatory subunit and a plausible binding site within that domain. The C-terminal half of the regulatory subunit also plays a crucial role in allowing valine binding, although the nature of this role requires further structural information on AHAS holoenzymes.

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ACKNOWLEDGMENTS We are greatly indebted to Gerhard Hu¨bner, Kay Tittmann, and many of their coworkers at Halle for the opportunity to collaborate with them, for kinetic measurements on AHAS II, and for valuable discussions. Many of our students in Beer-Sheva not listed as coauthors also contributed in one way or another to the work described here. Dr. Ahuva Bar-Ilan and Vitaly Balan each designed and prepared a variety of expression constructs, prepared various mutants, and played active roles in developing ideas about AHAS in the course of their Ph.D. work. Hagay Shmuely initiated and carried out mutagenesis of V375 and V461 in the course of his M.Sc. Judith Zohar, Udi Qimron, and Valery Vinogradov each prepared and carried out preliminary characterization of at least one mutant protein in the course of undergraduate senior projects; Udi first suggested making mutants at M250. Michael Vinogradov, whose biophysical work on subunit assembly we have not described for lack of space, actively contributed ideas on the regulatory subunits. We also thank Dr. Jerry Eichler for reading the ms and suggesting improvements. Finally, we are grateful for support of this work by research grants from the Israel Science Foundation (grants 243/98 and 660/01), a grant from the Yeshaya Horwitz Fund administered by B. G. Negev Technologies, and seed grants from the Vice President for Research and Development at BGU. REFERENCES 1. 2.

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JB Green. Pyruvate decarboxylase is like acetolactate synthase (ILV2) and not like the pyruvate dehydrogenase E1 subunit. FEBS Lett 246:1–5, 1989. TL Bowen, J Union, DL Tumbula, WB Whitman. Cloning and phylogenetic analysis of the genes encoding acetohydroxyacid synthase from the archaeon Methanococcus aeolicus. Gene 188:77–84, 1997. Y-Y Chang, JE Cronan Jr. Common ancestry of Escherichia coli pyruvate oxidase and the acetohydroxy acid synthases of the branched-chain amino acid biosynthetic pathway. J Bacteriol 170:3937–3945, 1988. YA Muller, G Schumacher, R Rudolph, GE Schulz. The refined structures of a stabilized mutant and of wild-type pyruvate oxidase from Lactobacillus plantarum. J Mol Biol 237:315–335, 1994. M Ibdah, A Bar-Ilan, O Livnah, JV Schloss, Z Barak, DM Chipman. Homology modeling of the structure of bacterial acetohydroxy acid synthase and examination of the active site by site-directed mutagenesis. Biochemistry 35:16282–16291, 1996. HE Umbarger. Biosynthesis of the branched-chain amino acids. In: FC Neidhardt, JL Ingraham, BL Low, B Magasanik, M Schaechter, HE Umbarger, eds. Escherichia coli and Salmonella typhimurium. Cellular and Molecular Biology. Washington, DC: ASM Press, 1996, pp. 442–457.

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Z Barak, DM Chipman, N Gollop. Physiological implications of the specificity of acetohydroxy acid synthase isozymes of enteric bacteria. J Bacteriol 169: 3750–3756, 1987. S Epelbaum, RA LaRossa, TK VanDyk, T Elkayam, DM Chipman, Z Barak. Branched-chain amino acid biosynthesis in Salmonella typhimurium: a quantitative analysis. J Bacteriol 180:4056–4067, 1998. N Gollop, B Damri, DM Chipman, Z Barak. Physiological implications of the substrate specificities of acetohydroxy acid synthases from varied organisms. J Bacteriol 172:3444–3449, 1990. JV Schloss, LM Ciskanik, EF Pai, C Thorpe. Acetolactate synthase: a deviant flavoprotein. In: B Curti, S Rochi, eds. Flavins and Flavoproteins. Berlin: Walter de Gruyter, 1991, pp. 907–914. SS Pang, RG Duggleby, LW Guddat. Crystal structure of yeast acetohydroxyacid synthase: a target for herbicidal inhibitors. J Mol Biol 317:249–262, 2002. A Bar-Ilan, V Balan, K Tittmann, R Golbik, M Vyazmensky, G Hubner, Z Barak, DM Chipman. Binding and activation of thiamin diphosphate in acetohydroxyacid synthase. Biochemistry 40:11946–11954, 2001. RG Duggleby, SS Pang. Acetohydroxyacid synthase. J Biochem Mol Biol 33:1– 36, 2000. D Chipman, Z Barak, JV Schloss. Biosynthesis of 2-aceto-2-hydroxy acids: acetolactate synthases and acetohydroxyacid synthases. Biochim Biophys Acta 1385:401–419, 1998. H Shmuely. Effect of the size of two amino acids in the active site of acetohydroxyacid synthase. MSc dissertation, Ben-Gurion University, Beer-Sheva, Israel, 1999. N Gollop, B Damri, Z Barak, DM Chipman. Kinetics and mechanism of acetohydroxy acid synthase isozyme III from Escherichia coli. Biochemistry 28: 6310–6317, 1989. B Mazur, SC Falco. The development of herbicide resistant crops. Annu Rev Plant Physiol 40:441–470, 1989. J Guardiola, M DeFelice, A Lamberti, M Iaccarino. The acetolactate synthase isoenzymes of Escherichia coli K-12. Mol Gen Genet 156:17–25, 1977. C Sella, O Weinstock, Z Barak, DM Chipman. Subunit association in acetohydroxy acid synthase isozyme III. J Bacteriol 175:5339–5343, 1993. O Weinstock, C Sella, DM Chipman, Z Barak. Properties of subcloned subunits of bacterial acetohydroxy acid synthases. J Bacteriol 174:5560–5566, 1992. M Vyazmensky, C Sella, Z Barak, DM Chipman. Isolation and characterization of subunits of acetohydroxy acid synthase isozyme III and reconstitution of the holoenzyme. Biochemistry 35:10339–10346, 1996. S Mendel, T Elkayam, C Sella, M Vyazmensky, DM Chipman, Z Barak. Acetohydroxyacid synthase: a proposed structure for regulatory subunits supported by evidence from mutagenesis. J Mol Biol 307:465–477, 2001. R Al Rabiee, J Lee Edward, A Grant Gregory. The mechanism of velocitymodulated allosteric regulation in D-3-phosphoglycerate dehydrogenase: crosslinking adjacent regulatory domains with engineered disulfides mimics effector binding. J Biol Chem 271:13013–13017, 1996.

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24. DJ Schuller, GA Grant, LJ Banaszak. The allosteric ligand site in the V-maxtype cooperative enzyme phosphoglycerate dehydrogenase. Nature Struct Biol 2:69–76, 1995. 25. MD Feese, HR Faber, CE Bystrom, DW Pettigrew, SJ Remington. Glycerol kinase from Escherichia coli and an Ala65!Thr mutant: the crystal structures reveal conformational changes with implications for allosteric regulation. Structure 6:1407–1418, 1998. 26. DM Chipman, B Shaanan. The ACT domain family. Current Opinion in Structural Biology 11:694–700, 2001. 27. L Aravind, EV Koonin. Gleaning non-trivial structural, functional and evolutionary information about proteins by iterative database searches. J Mol Biol 287:1023–1040, 1999. 28. S Engel, M Vyazmensky, Z Barak, DM Chipman, JC Merchuk. Determination of the dissociation constant of valine from acetohydroxy acid synthase by equilibrium partition in an aqueous two-phase system. J Chromatogr B 743: 225–229, 2000. 29. JMT Tse, JV Schloss. The oxygenase reaction of acetolactate synthase. Biochemistry 32:10398–10403, 1993. 30. GA Grant, DJ Schuller, LJ Banaszak. A model for the regulation of D-3phosphoglycerate dehydrogenase, a V-max-type allosteric enzyme. Protein Sci 5:34–41, 1996. 31. PJ Kraulis. MOLSCRIPT: A program to produce both detailed and schematic plots of protein structures. J Applied Crystallography 24:946–950, 1991.

15 Structure and Properties of Acetohydroxyacid Synthase Ronald G. Duggleby, Luke W. Guddat, and Siew Siew Pang The University of Queensland, Brisbane, Australia

I. INTRODUCTION A. Branched-Chain Amino Acid Biosynthesis L-Valine, L-leucine,

and L-isoleucine are synthesized by a common pathway in microorganisms and plants. The common precursor for these amino acids is pyruvate, while L-isoleucine biosynthesis requires a second precursor, 2-ketobutyrate. One notable feature of this pathway is the employment of parallel steps leading to the formation of L-valine and L-isoleucine. These parallel steps involve four enzymes, namely, acetohydroxyacid synthase (AHAS; EC 4.1.3.18), ketol-acid reductoisomerase, dihydroxyacid dehydratase, and a transaminase, each of which is capable of catalyzing two similar reactions using two alternative substrates. AHAS (1) catalyzes the first of these parallel steps and produces either 2-acetolactate or 2-aceto2-hydroxybutyrate (Fig. 1a). Each of the products is then converted further in three reactions to give L-valine and L-isoleucine, respectively. For L-leucine biosynthesis, four additional enzymes are required, using the L-valine precursor 2-ketoisovalerate and acetyl CoA as the starting point for synthesis. 251

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Figure 1 Reactions catalyzed by (a) and herbicidal inhibitors of (b) AHAS.

B. Cofactors The reactions catalyzed by AHAS involve an initial decarboxylation of pyruvate; as expected, the enzyme requires thiamine diphosphate (TDP) as an essential cofactor. The bound hydroxyethyl-TDP intermediate condenses with either a second pyruvate to form acetolactate or 2-ketobutyrate to give acetohydroxybutyrate. In common with other TDP-dependent enzymes, AHAS requires a divalent metal ion that anchors TDP to the protein. An unexpected feature of AHAS is the additional requirement for FAD. The reaction catalyzed involves no net oxidation, so the requirement for FAD is clearly not for any redox chemistry. Some bacteria, such as Klebsiella pneumoniae, contain a second form of the enzyme that is independent of FAD (2)

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and is specific for acetolactate formation. The existence of this enzyme demonstrates that there is no chemical feature of the AHAS reaction that demands the participation of FAD. Indeed, this FAD-independent form has a turnover number that is 5- to 10-fold higher than is usually observed for the FAD-dependent enzyme. C. Distribution and Herbicides The branched-chain amino acid biosynthetic pathway and the enzymes comprising it are widely distributed in nature but not found in higher animals. A reported human AHAS (3) has no AHAS activity (4) and now appears to be a 2-hydroxyphytanoyl CoA lyase (5). Eukaryotic AHAS is localized in either mitochondria (fungi) or chloroplasts (plants). Due to the absence of this pathway in animals, these enzymes have attracted attention as potential targets for pesticides and antibiotics (6). This potential has been fulfilled for several families of commercial herbicides that act by inhibiting plant AHAS. Typical examples are shown in Figure 1b; chlorsulfuron is one of the sulfonylurea family (7) devised by Du Pont, while imazapyr is one of Cyanamid’s imidazolinones (8). These are very effective herbicides that appear to be very safe; in acute doses they are less toxic to rats than is common salt. D. Sources of AHAS Due to the interest in AHAS as a herbicide target, a considerable amount of the published work has focused on the plant enzyme. For the most part these studies have used crude plant extracts or partially purified enzyme only. The development of heterologous-expression systems to overproduce plant AHAS (9–11) have allowed more detailed characterization of the enzyme. In our laboratory we have purified and characterized recombinant Arabidopsis thaliana AHAS (12) as well as isoenzyme II from Escherichia coli (13) and yeast AHAS (14). It is the yeast enzyme that will be the main focus of this chapter. E. AHAS Subunits Bacterial AHAS is composed of two types of subunit (15). The larger subunit (f60 kDa) has sequence homology to several other TDP-dependent enzymes, such as pyruvate decarboxylase (PDC), benzoylformate decarboxylase (BFDC), and pyruvate oxidase (POX). It contains all of the catalytic machinery and is usually active alone (12,14,16). The smaller subunit (9–17 kDa) plays a regulatory role, stimulating the activity of the catalytic subunit

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and mediating inhibition by one or more of the branched-chain amino acids, usually valine (17). The FAD-independent enzyme contains no regulatory subunit and is not inhibited by branched-chain amino acids. Until recently, there were several tantalizing clues suggesting the existence of a regulatory subunit of eukaryotic AHAS (18) but no definite proof. For example, the eukaryotic enzyme in crude extracts is sensitive to branched-chain amino acid inhibition, while the catalytic subunit gene expressed in E. coli gives an enzyme that is totally insensitive to these compounds (19). We therefore set about trying to identify possible eukaryotic regulatory subunits and chose yeast for these studies. F. Why Yeast? There were several reasons that led us to choose yeast for this search. First, we already had a clone of ilv2, the yeast catalytic subunit. The gene had been expressed in E. coli (20), and while the enzyme was active it showed no inhibition by valine (14). We had also identified a yeast DNA sequence that seemed to meet the necessary criteria to encode a regulatory subunit (14,18). This ilv6 gene has sequence homology with bacterial regulatory subunit genes. It encodes a protein with what appears to be an N-terminal mitochondrial transit peptide, necessary because it was known that the fungal enzyme is located in mitochondria. DNA sequence analysis identified expected regions, such as the GCN4 transcription activator-binding sequence that controls a number of amino acid biosynthetic pathways in yeast (21). And finally, AHAS in yeast cell extracts of an ilv6 knockout mutant had been shown to be insensitive to feedback regulation (22).

II. ENZYMATIC CHARACTERIZATION A. Expression of Yeast AHAS in E. coli To obtain large quantities of enzyme for kinetic and structural characterization, the yeast catalytic subunit gene (ilv2) and the putative regulatory subunit gene (ilv6) were cloned and overexpressed in E. coli (14). Both of the genes were cloned into pET30(+) expression vectors, under the control of the strong T7 promoter. The cloning procedure also introduced N-terminal hexahistidine tags on the recombinant proteins to allow for easy and rapid purification by immobilized-metal affinity chromatography of the otherwise labile proteins. The initial plasmid construct containing the full-length sequence of ilv2 produced an expressed protein that is found mostly in the insoluble fraction of bacterial lysates [14,20]. Deletion of 57 or 81 amino acids from the N-terminal mitochondrial transit peptide produced soluble and ac-


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tive enzyme. Similarly, partial removal of the mitochondrial transit peptide is important for the expressed regulatory subunit to stay in solution (14). B. Conditions for In Vitro Yeast AHAS Reconstitution If the ilv6 gene product is to function as the regulatory subunit of yeast AHAS, we anticipated that it would interact with the catalytic subunit to give a reconstituted activity with properties similar to those observed in vivo. Studies of E. coli isozymes II (13) and III (17) showed that full AHAS activity can be reconstituted in vitro by combining the catalytic and regulatory subunits that are expressed and purified separately. Reconstituted-AHAS activity is characterized by enhanced specific activity in both enzymes and by the acquisition of valine feedback inhibition in the case of AHAS III. When yeast AHAS reconstitution experiments were conducted in low-ionic-strength buffers (50 mM potassium phosphate, pH 6.51–7.8), the addition of the ilv6 protein to the catalytic subunit, in the presence or absence of valine, resulted in no observable change in the enzymatic activity. Several papers on fungal AHAS published around 1970 offered an explanation for these observations. The sensitivity to valine inhibition of AHAS in yeast cells is extremely labile, with the regulation of activity being lost rapidly in crude extracts when the enzyme was solubilized from cells or intact mitochondria (23). Similar results were observed with the mitochondrial-localized Neurospora crassa AHAS activity (24–26). A later report (27) showed that the high AHAS specific activity and valine sensitivity in intact N. crassa mitochondria could be preserved if the enzyme was extracted using buffer containing 1 M phosphate at neutral pH. Accordingly the effect of phosphate concentration on the reconstitution of yeast AHAS from its subunits was tested. At the optimal phosphate concentration of about 1 M at neutral pH, the AHAS activity is stimulated by as much as sixfold (Fig. 2). With the addition of 5 mM valine, the increased activity is abolished, indicating that the reconstituted enzyme is regulated by end-product feedback inhibition. The activity of the catalytic subunit alone is mostly unaffected by the presence of phosphate (0.04–1.5 M), in the presence or absence of valine (14). These data provided the first biochemical evidence that eukaryotic AHAS, similar to the bacterial enzyme, is composed of two different subunits and that the yeast ilv6 gene encodes the regulatory subunit. The reconstitution of yeast AHAS was tested further with a variety of salts. Potassium phosphate is routinely used in our yeast AHAS reconstitution assays. This salt can be fully replaced by similar concentrations of sodium phosphate and, to a lesser extent, by sulfate salts of potassium and sodium. Further experiments were conducted to investigate the role of high phosphate concentration in yeast AHAS reconstitution. Using tryptophan spectro-

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Figure 2 Effect of potassium phosphate concentration on yeast AHAS. Catalytic subunit alone (.); reconstituted enzyme (n); reconstituted enzyme with 5 mM valine (E).

fluorimetry and size-exclusion chromatography, the high phosphate concentration was shown to be required for the physical interaction of the catalytic and regulatory subunits; no interaction is observed under low-phosphate conditions (28). The requirement for high phosphate concentrations has only been reported with the mitochondrial-localized fungal AHAS (14,27) and is not required for the integrity of bacterial (13,17) and plant (29,30) AHAS. This unusual condition may mimic the in vivo mitochondrial environment in which fungal AHAS resides. C. Regulation of Yeast AHAS Yeast AHAS reconstituted by combining the separate subunits, like the native enzymatic activity, is regulated by an end-product feedback mechanism. Out of the three branched-chain amino acids, valine is the most potent inhibitor, with an apparent Ki value of 73 lM (Fig. 3a). The inhibition by valine is partial; at a saturating valine concentration of 5 mM, a residual AHAS activity of about 15% remains. The inhibitory effects of the other two branched-chain amino acids are minimal; isoleucine results in a slight inhibition at the highest concentration tested (5 mM), while this concentration of leucine has no effect on the reconstituted activity (Fig. 3a).


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Figure 3 Regulation of reconstituted yeast AHAS. (a) Inhibition by branchedchain amino acids. (b) Activation by ATP of the enzyme inhibited by valine.

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In addition to end-product feedback regulation, yeast AHAS is regulated by MgATP (28). The regulation by ATP was first observed with the AHAS activity in intact mitochondria of N. crassa (26) and Euglena gracilis (31) and has never been described for enzymes from bacteria or higher plants. In yeast AHAS, MgATP activates the valine-inhibited enzyme (Fig. 3b). Even in the absence of valine, the reconstituted enzyme appears to be slightly stimulated by MgATP, resulting in an activity increase of about 20%. Since MgATP has no effect on the catalytic subunit alone, the activation appears to be imposed upon the enzyme via the regulatory subunit. Indeed we have shown by circular dichroism that ATP, in the presence of Mg2+, binds directly to the regulatory subunit with a Kd value of 0.24 mM (28). The location of the MgATP-binding site on the protein is not known, since the regulatory subunit does not contain any of the known ATP-binding consensus sequences described in the Prosite database (http://www.expasy.ch/ prosite/). Analysis of the sequence of ilv6 and other fungal AHAS homologous sequences (presumed regulatory subunits) reveals an extra segment of about 50 amino acids in the middle of the sequence that is not present in the bacterial (14) and plant (29,30) regulatory subunits. Since ATP does not activate the bacterial and plant enzymes, these extra 50 residues in the yeast subunit might constitute part or all of the MgATP-binding site. We are currently carrying out mutagenesis studies to identify residues that are important for MgATP and valine binding to yeast AHAS. The regulation of yeast AHAS by valine and MgATP is highly specific (28). As mentioned earlier, valine cannot be replaced by leucine or isoleucine; other valine analog tested and shown to be ineffective are valineamide, N-methylvaline, and N-acetylvaline. Similarly, the activation by ATP cannot be mimicked by ADP, AMP, GTP, UTP, CTP, and N6-ethenoATP. The only effective ATP analog tested is AMP-PNP, and this rules out the possibility that the activation is due to autophosphorylation of the enzyme. Escherichia coli and Salmonella typhimurum have at least three functional isozymes of AHAS. The presence of different isozymes, each with distinct kinetic properties, are thought to allow the enterobacteria to adapt and survive under the varying conditions in their natural environment (32,33). Among the isozymes, AHAS II is uniquely valine insensitive. Wild-type E. coli strain K-12 is a laboratory isolate and lacks AHAS II activity due to a frameshift mutation in the catalytic subunit (34). As a result, this strain of E. coli is unable to grow in minimal medium supplemented with valine, due to isoleucine starvation. Yeast has a single AHAS gene and, in contrast to wild-type E. coli, has no valine-insensitive AHAS isozyme. However, the growth of yeast cells in minimal medium is not inhibited by valine (35). This is not surprising, since valine inhibition is partial (Fig. 3a) and the valine-inhibited enzyme can be activated by ATP at the concentrations that are found in actively growing yeast cells (36). Thus, instead of having multiple isozymes,


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yeast cells may rely upon partial inhibition combined with ATP activation to achieve the same result as having a valine-insensitive isozyme in E. coli. III. STRUCTURE OF AHAS A. Crystallization of Yeast AHAS The catalytic subunit of yeast AHAS has been crystallized by the hangingdrop vapor diffusion method in the presence of all cofactors (TDP, MgCl2, and FAD). The crystals are formed under slightly acidic conditions in the presence of potassium phosphate, PEG 4000, and ammonium acetate (37). Potassium phosphate is important for stabilizing the enzyme (14), while the other two reagents are precipitants commonly used for protein crystallization. The AHAS crystals are yellow in color, due to bound FAD, and grow to a typical size of 0.2 0.2 0.5 mm. X-ray diffraction data to 2.6 A˚ were collected at the synchrotron facility at Advance Proton Source, Chicago. The data were phased by molecular replacement using selected regions of the crystal structure of BFDC as the search model (37). The structure was refined to Rfactor and Rfree values of 0.188 and 0.219, respectively. B. General Features of the Three-Dimensional Structure of Yeast AHAS The catalytic subunit of yeast AHAS crystallizes as a dimer (38), which is the same subunit structure observed in solution. The subunits are tightly associated, forming numerous noncovalent interactions across the dimer interface. The surface area buried by this interface is about 5000 A˚2. In each monomer, there are three domains, designated a, h, and g (Fig. 4). The surfaces of the a-domains and g-domains in each monomer form the subunit interface. The h-domains are distal to each other and play only a minor role in stabilizing the dimer interface. The a-domain consists of amino acid residues 85–269 in the polypeptide chain, the h-domain comprises residues 281–458, and the g-domain includes residues 473–643. Interestingly, all three domains have a similar arrangement of secondary structure elements, consisting of a central six-stranded parallel h-pleated sheet flanked by six to eight a-helices. The a-domain is slightly larger than the h- and g-domains. This is due to the presence of a two-stranded antiparallel h-sheet that links the amino- and carboxy-terminal ends of the domain. In our structure of yeast AHAS, the polypeptide sections that link the domains are not well resolved. In particular, no electron density is visible for some of the connector (residues 271–279) between the a-domain and the h-domain in one of the monomers. Other connector segments are hallmarked by high average isotropic temperature factors, for some atoms in excess of 100 A˚2. In general, the polypeptide segments within the domains

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Figure 4 Structure of the catalytic subunit monomer of yeast AHAS.

are well resolved, and individual atoms have temperature factors less than 50 A˚2. One exception is a segment of polypeptide that includes amino acid residues 580–595. In one of the subunits this section is completely disordered, while it is traceable in the other subunit. We hypothesize that the structure may be ordered due to close crystal packing contacts. The amino acid residues in this region are of critical importance since they form part of the active site of the enzyme and mutations in this region result in insensitivity to herbicidal inhibitor compounds. As mentioned previously, an open reading frame discovered in yeast has been confirmed to be an AHAS regulatory subunit. Reconstitution with the yeast catalytic subunit results in a 7- to 10-fold increase in activity and confers upon it sensitivity to valine inhibition (14). It is interesting to speculate on the


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location of the regulatory subunit relative to the catalytic subunit. Based on the increase in activity, we expect that at least some portion of the regulatory subunit would form a close association with the active site, possibly helping to sequester the substrates during catalysis. We are currently working on obtaining crystals of the complex between the catalytic and regulatory subunits. This will allow us to understand how the complete yeast AHAS machinery is assembled. Comparison of the sequence of the regulatory subunit of AHAS with that of the regulatory domain of threonine deaminase suggests that they could have a very similar overall three-dimensional fold (30), although there is presently only limited experimental data to support this hypothesis (39). C. Comparison with Other Thiamine Diphosphate– Dependent Enzymes The overall structure of the catalytic subunit of yeast AHAS is very similar that of other TDP-dependent enzymes, such as BFDC and POX. Evidence for this comes from calculation of the root mean square deviation (rmsd) values when equivalent core a carbon atoms within the individual domains of the three proteins are superimposed (Fig. 5). Typically, the rmsd values vary between 1.2 A˚ and 1.7 A˚, indicative of a high degree of structural similarity. The largest variation in structure when comparing the three enzymes is the location of the h-domain relative to the a- and g-domains. The h-domains of BFDC and AHAS are only slightly misaligned relative to the a- and g-domains. However, the orientation of the h-domain in POX is considerably different when compared to the other enzymes. Inspection of the structures of the connectors that lead in and out of the h-domain shows some interesting differences. In AHAS the connection between the a- and h-domains is longer by eight amino acid residues when compared to BFDC and by six amino acid residues when compared to POX. The effect of this difference is that the connector tends to bulge out in yeast AHAS relative to the other two enzymes. It may be that the long helical connection between the h- and g-domains of POX, not observed in the other two enzymes, accounts for the different orientations of the h-domain. The connection between the h-domain and g-domains is different in all three enzymes. In terms of the number of amino acid residues, BFDC has the shortest connection between these two domains by using a polyproline type II helix [40]. In AHAS, a small segment of a-helical structure is found as part of the carboxy-terminal end of the h-domain, but the remainder of the connection is extended random structure. POX, with the longest connection in terms of the number of amino acid residues between the two domains, adopts a completely a-helical structure.

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Figure 6 The channel in the h-domain of yeast AHAS containing FAD.

D. The Active Sites and Location of the Cofactors in AHAS Each AHAS has two active sites, and, by virtue of symmetry, they are found on opposite faces of the molecule. Each active site is bordered by amino acids from both monomers. The two organic cofactors bound to AHAS are TDP and FAD. Except for a single van der Waals interaction between the C8 methyl group of the flavin ring and the side chain of a phenylalanine residue from the a-domain of the second monomer, the FAD is bound almost exclusively to amino acid residues in the h-domain and lies in a channel formed by this domain (Fig. 6). TDP is anchored through a magnesium ion to the g-domain, but it also has significant interactions with Glu139 and three other amino acids from the a-domain of the second monomer. The binding of the two organic cofactors to AHAS is similar, in the sense that the attachment sites are formed by loops located at the ends of the central h-sheet structures in the three domains.

Figure 5 Overlay of Ca atoms of TDP-dependent enzymes. (a) Yeast AHAS (light shade) and POX (dark shade). (b) yeast AHAS (light shade) and BFDC (dark shade).

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1. TDP TDP is centrally located in the active site spanning the two monomers (Fig. 7). The thiazolium and pyrimidine rings of this cofactor are held to the enzyme in a V-shaped conformation by van der Waals interactions to Met525, Met555, Tyr113, Gly523, and Ala551 and by two hydrogen bonds, one between Glu139 and N1V of the pyrimidine ring and the other between the backbone oxygen atom of Gly523 and the 4V-amino/imino group. The diphosphate tail is anchored to the protein through interactions to a magnesium ion and several water molecules. There are six ligands to the magnesium ion: two phosphate oxygen atoms, two protein ligands (Asp550 and Asn557), and two water molecules. The overall geometry around the metal is a distorted octahedron. 2. FAD FAD is bound to the enzyme in an extended conformation and is most closely associated with the h-domain (Fig. 6). In all, FAD forms 12 hydrogen bonds and at least 22 van der Waals interactions with the protein. We have recently measured that yeast AHAS has a Km of 27 nM for FAD, much lower than the

Figure 7 Two views of the residues contacting TDP in yeast AHAS. Amino acids from different monomers are shown in different shades of gray.


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previously reported value of 0.3 lM (20). Given the large number of noncovalent interactions, our value for Km would seem to be more reasonable. FAD is an essential cofactor for AHAS activity. Since the reaction does not involve any electron transfer, the function of the flavin cofactor in AHAS is unclear and has been suggested to be solely structural. The noncatalytic role of FAD is further supported by the existence in some bacteria of a catabolic enzyme (acetolactate synthase) that catalyzes an AHAS reaction (acetolactate production) and is FAD free (2). Based on the sequence and catalytic similarities with the ThDP- and FAD-dependent enzyme POX, AHAS has been proposed to have evolved from a POX-like ancestral enzyme (41). It has also been suggested that FAD in AHAS is a vestigial remnant, which provides an explanation for the apparent structural role of the cofactor. Overall, the structure of the catalytic subunit of yeast AHAS does not permit any definite conclusion to be drawn about the role of FAD, other than structural. By comparing the threedimensional structures, particularly the active sites of AHAS and POX, we hope to gain a better understanding of why FAD is redox active in the latter enzyme but not in the former. In both POX and AHAS, FAD is bound in a similar fashion and with most of the numerous interactions between the cofactor and the protein conserved. The main differences lie in the conformation and position of the flavin ring (Fig. 8). In POX, even in the absence of substrate, the isoalloxazine ring of FAD is bent by 15j across the N5–N10 axis, whereas in AHAS it is planar. The explanation for the bend in POX is that it promotes the reduction of FAD, facilitating electron transfer during the reaction. The crystal structure of POX does not allow definitive conclusions to be drawn about the route through which electrons are transferred from the reaction intermediate to FAD (42). The positions of the flavin N5 atom and the active center (C2 of TDP) are too far apart to allow direct transfer without significant movement of the cofactors during reaction. Several indirect routes have been proposed and are yet to be proven experimentally. In AHAS, the N5 atom of FAD is also a large distance (13.3 A˚) away from the active center. In addition, when superimposed on the active site of POX, the flavin ring in AHAS is pointing away from the active site. The planar conformation, position, and orientation of FAD in AHAS may all contribute to its inability to participate in electron transfer. IV. HERBICIDE-BINDING SITE A. Herbicide-Resistance Mutations A number of AHAS mutations have been found that result in reduced sensitivity to herbicidal inhibitors (1). These have been discovered in field

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Figure 8 Comparison of the locations and conformations of cofactors in yeast AHAS and POX. TDP is superimposed, while FAD is shown for POX (light shade, ball and stick) and AHAS (dark shade, stick).


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isolates and by random or targeted mutagenesis of AHAS genes. The most extensive series of these are those reported for yeast AHAS (43), where mutation at 10 sites each results in resistance to the sulfonylurea sulfometuron methyl. At some sites, almost any amino acid substitution results in resistance, while at others only a few substitutions are tolerated. Studies of AHAS from plants and bacteria have identified other herbicide-resistance mutation

Figure 9 The herbicide-binding site of yeast AHAS. (a) Locations of sites in the AHAS amino acid sequence where mutation results in herbicide resistance. (b) Proposed interaction of AHAS and the herbicidal inhibitor, imazapyr. Amino acid residues at herbicide-resistance sites are labeled and shown as CPK models. Residues with and without the prime symbol (V) are derived from different monomers. (Adapted from Ref. 38, Fig. 9.)

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sites and also confirmed some of those observed for the yeast enzyme. The location of 15 sites are shown schematically in Figure 9a, which reveals that the mutations are spread across the entire sequence and include residues from all three domains of the protein. The natural amino acid at most of these sites is strongly conserved across many species, suggesting that these residues play important roles. However, the fact that organisms carrying these mutations are viable but herbicide resistant shows that the variants must be active. Indeed, detailed in vitro studies of some variants (44–47) have shown that they have no major alterations in activity or the kinetics toward substrate or cofactors. Thus, the role of these residues and the selection forces involved in their maintenance are unanswered questions. B. Structure of the Herbicide-Binding Site Mapping of the herbicide-resistance sites on to the three-dimensional structure of yeast AHAS has allowed us to delineate the herbicide-binding site (38). This is shown in Figure 9b, with a possible position for imazapyr (Fig. 1b) obtained by docking calculations (48). Several features are evident from this structure. First, the herbicide-binding site is located at the subunit interface and is bounded by residues from the a-domain of one monomer and the hand g-domains of the second monomer. Second, the site is near to the active site and in close proximity to both TDP and FAD. Thus, it is easy to imagine how herbicides might block access by substrate, resulting in inhibition. Third, the site is rather large compared to the size of imazapyr and the position of the herbicide is quite distant from some residues, especially those in the h- and g-domains. One possible explanation is that imazapyr is not correctly docked and should be in another region of the site. However, no matter how imazapyr is positioned, it could not interact simultaneously with all of these residues without some structural alteration of the protein. The published literature on the effects of mutation at each site on sensitivity to particular herbicides is relatively sparse, and in only a few cases (44–47) has the specificity toward a range of different herbicides been determined. Therefore, we have recently undertaken a systematic program of mutagenesis of residues in the proposed herbicide-binding site of yeast AHAS and characterization of the inhibition by a series of sulfonylurea and imidazolinone herbicides. For example, we have now shown that mutation at G116, A200, K251, M354, W586, and F590 all result in imazapyr resistance and that changes to A117, L119, P192, S194, and V583 have little effect. Curiously, mutation of D379 results in an enzyme that is nearly 50-fold more sensitive to imazapyr inhibition, and mutation of G657 also increases imazapyr sensitivity. For chlorsulfuron (Fig. 1b), all of the mutations that we have tested to date (at G116, A117, P192, A200, K251, M354, D379, V583,


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W586, and F590) result in resistance. These results strongly suggest that both types of herbicide interact with residues from all three domains, implying that the rather open structure shown in Fig. 9b must be more closed when herbicides are bound. It is relevant that the herbicide-resistance residues in the g-domain are within the disordered region (residues 580–595) described earlier. Thus, it seems probable that this region forms a mobile loop that closes in the active enzyme or upon binding of herbicides. Further active-site closure may also involve movement of the h-domain. C. Natural Role of the Herbicide-Binding Site As mentioned earlier, the presence of FAD in AHAS has been proposed to be an evolutionary remnant of an ancestral, POX-like enzyme (41). Extending this argument, it has been suggested (49) that the herbicide-binding site is a relic of the binding site for the quinone substrate of this ancestral enzyme. This hypothesis begs the question of why the site has been maintained so that AHAS from plants, fungi, and bacteria are all inhibited by herbicides. It was noted earlier that residues surrounding the herbicide-binding site show strong conservation across species, suggesting that this region plays some positive role. Here we wish to propose an entirely speculative idea on this matter. The first half of the reaction catalyzed by AHAS is identical to that of some other TDP-dependent enzymes, such as PDC, POX, and the E1 component of pyruvate dehydrogenase. After decarboxylation, the resonating carbanion/enamine of the bound hydroxyethyl intermediate will be formed, and it is essential to protect this from protonation by solvent and subsequent release as acetaldehyde. However, the active site must also accept the second substrate (pyruvate or 2-ketobutyrate), which then reacts with the intermediate to form the acetohydroxyacid product. It is difficult to imagine how the active site could open to admit the second substrate without allowing the entry of solvent protons. Our suggestion is that the active site does not open midway though the catalytic cycle. Rather, the herbicide-binding site acts as a ‘‘waiting room’’ that contains the second substrate, ready to react with carbanion/enamine once it is formed. This site would therefore be vital for AHAS to act efficiently and would explain the conservation of the herbicide-binding site through evolution and across species. Although there is scant experimental evidence to support this hypothesis, it does agree with the suggestion that herbicides compete with the binding of the second substrate (50).

V. FUTURE STRUCTURES The AHAS structure described earlier is of the catalytic subunit only, with no bound ligands. The position of imazapyr was deduced from docking calcu-

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lations but has not been verified experimentally. Clearly, determination of a structure with a bound imidazolinone or sulfonylurea herbicide would be informative, and we are actively pursuing this goal. Solving the structure of yeast AHAS containing the sulfonylurea herbicide chlorimuron ethyl is now nearing completion. The regulatory subunit greatly enhances the activity of the catalytic subunit, but the mechanism of this activation is unknown at the molecular level. Two partial models for the structure of the regulatory subunit have been proposed (30,51), but each has, at best, only limited experimental support (39,51). We have tried to crystallize the regulatory subunit and the complex formed between the catalytic and regulatory subunits but have not yet been successful. A complex containing the inhibitor valine would also be very interesting, as would one with the activator MgATP. As mentioned previously, MgATP activation has been observed for the fungal enzyme only and is expected to involve interactions with a 50-residue protein segment that is found only in the fungal regulatory subunit. Despite now knowing the structure of AHAS, the role of FAD remains elusive. One way to explore this role is to compare our structure with that of the bacterial FAD-independent enzyme. We have had some success in this area and have obtained crystals of the K. pneumoniae enzyme (52) that diffract to 2.1 A˚. Solving this structure is now well under way. ACKNOWLEDGMENTS This work was supported by grants A09937067 and A00105313 from the Australian Research Council. The use of the BioCARS, Argonne National Laboratory, Chicago, Illinois, was supported by the Australian Synchrotron Research Program. REFERENCES 1. 2. 3.

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16 Exploring the Substrate Specificity of Benzoylformate Decarboxylase, Pyruvate Decarboxylase, and Benzaldehyde Lyase Petra Siegert and Martina Pohl Heinrich-Heine-Universita¨t Du¨sseldorf, Ju¨lich, Germany Malea M. Kneen, Irina D. Pogozheva, George L. Kenyon, and Michael J. McLeish University of Michigan, Ann Arbor, Michigan, U.S.A.

I. INTRODUCTION Benzoylformate decarboxylase (BFD; EC 4.1.1.7) catalyzes the nonoxidative cleavage of benzoylformate to benzaldehyde and carbon dioxide (Fig. 1). The mandelate pathway, in which BFD is found, allows several closely related microorganisms to use R-mandelate as a sole carbon source by converting it to benzoic acid, which is then metabolized by the h-ketoadipate pathway and the citric acid cycle (1–6). This pathway is believed to have evolved relatively recently, and the members of the pathway have attracted interest as models for the studies of enzyme functional evolution (7). Under anaerobic conditions, pyruvate, the product of glycolysis in yeast, undergoes nonoxidative cleavage to acetaldehyde and carbon dioxide 275

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Figure 1 Reactions catalyzed by benzoylformate decarboxylase, pyruvate decarboxylase, and benzaldehyde lyase.

(Fig. 1). The reaction is catalyzed by pyruvate decarboxylase (PDC, E.C. 4.1.1.1), and the acetaldehyde is subsequently converted to ethanol by alcohol dehydrogenase. In addition to yeast, the enzyme is widely distributed in plants and is also found in other fungi and some bacteria, particularly in the obligatory fermentative bacterium Zymomonas mobilis (8–10). Both BFD and PDC are thiamine diphosphate (TDP)–dependent enzymes and, in the active form, are tetrameric. The X-ray structures of BFD (11), PDC from Z. mobilis (ZmPDC; Ref. 12) and PDC from S. cerevisiae (ScPDC; Refs. 13 and 14) are available. These show that, overall, the architectures of the two decarboxylases are quite similar. Each active site contains two histidine residues, although these arise from different regions of the polypeptide chain. In BFD (15), ZmPDC (16,17), and ScPDC (18), both histidines have been implicated in catalysis. In addition, in the active site, Ser26 of BFD is superimposed on Asp28 of PDC, a residue that has also been shown to be involved in catalysis (18,19). In contrast to the two decarboxylases, benzaldehyde lyase (BAL), another TDP-dependent enzyme, has undergone little investigation. Initially, benzaldehyde lyase was isolated from Pseudomonas fluorescens biovar I (20) and was found to cleave the a-hydroxy linkage in benzoin to give two molecules of benzaldehyde (Fig. 1). More recently, it has been cloned (21), expressed, and found some utility in enantioselective synthesis (22). In addition to BAL, several TDP-dependent enzymes, including BFD and PDC, have proved useful in chemoenzymatic syntheses (22–24). As a consequence, we have been interested in identifying residues affecting the substrate specificity of these three enzymes. Initial studies indicated that, for

Substrate Specificity of BFD, PDC, and BAL

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ZmPDC, Ile472 and Ile476 affect both the substrate specificity and the stereoselectivity of the carboligation reaction (25). Here we identify the analogous residues in BFD and describe the preliminary steps toward converting ZmPDC into a BFD, and vice versa. Further, we develop a model for the active site of BAL that permits us to suggest residues essential for substrate binding and catalysis. II. MATERIALS AND METHODS A. Construction, Expression, and Purification of His-Tagged PDC Variants WT and PDCI472A were available from a previous study (25). Other PDC point mutations were introduced by PCR using the overlap extension technique (26). All fragments were amplified using the following outer primers: PDC (bp 879) PDC (bp 1717)

5V-GGTGGACGGATATCCCTGATCC-3V(sense) 5V-AGTAAGCTTCTAGAGGAGCTTGTTAAC-3V (antisense)

The PCR-generated fragments were initially cloned into pUC18 and sequenced. Fragments (635 bp) carrying the desired mutation were excised using EcoRV and StuI and cloned into the similarly digested expression vector pPDC-His6 (25). The forward primers used in the mutagenesis were: PDCI476F: PDCI472A/I476F:

5 -V A C A C C A T C G A A G T T A T G t T C C A T GATGG-3V 5V-ACACCgcCGAAGTTATGtTCCATGATGG-3V

In both cases the mutated codons are underlined, with the lowercase letters indicating a base change from wild type. All PDC variants were expressed and purified, essentially as described previously (25). The only modification was that chromatography was carried out in 50 mM potassium phosphate, pH 6.5, containing 2 mM MgSO4 and 0.1 mM TDP, rather than the Mes buffer used in the earlier study. B. Construction, Expression, and Purification of the BFD Variants With the exception of BFD S26A and H70A, which were available from previous studies (15,27), the BFD mutants used here were all prepared using Pfu DNA polymerase and the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA). The single mutants were prepared using pBFDtrc

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(28) as the DNA template, while pBFDF464I served as the template for the construction of the double mutant. The forward primers used for the mutagenesis are shown here. Again the mutated codons are underlined, and the lowercase letters indicate a base change from wild type: A460I: 5V-ATGAACAACGGCACgTACGGTatcTTGCGATGG-3V F464I: 5V-GCGTTGCGATGGaTTGCaGGCGTTCTCGAA-3V H70Q: 5V- GCCGGCTTTCATTAACCTGCAgTCTGCTGCTGG-3V In addition to creating the appropriate mutations, the change in sequence for BFD A460I resulted in the loss of a Ban1 restriction site. For BFDF464I, an NaeI restriction site was lost and for H70Q a BsmI site was lost. Following mutagenesis, the template DNA was removed by treatment with Dpn1 and the remaining PCR products transformed into E. coli strain XL1-Blue (Stratagene, La Jolla, CA). Single colonies were picked and their DNA isolated and screened for the desired mutation by restriction analysis using the appropriate combination of restriction enzymes. For all mutants, the fidelity of the PCR amplification and the presence of the mutation were confirmed by sequencing. Plasmids containing the mutated BFD were selected and denoted pKKH70Q, pKKBFDA460I, pKKBFDF464I, and pKKBFDA460I/F464I, respectively. After IPTG-induced expression in E. coli SG13009, the cells were harvested and disrupted with glass beads. The cell-free extract was applied onto a Q-Sepharose FF-column equilibrated with buffer A (50 mM potassium phosphate, pH 6.0, 0.1 mM TDP, 2 mM MgSO4, and 150 mM KCl). After elution of nonbound proteins with buffer A, a linear gradient of 200 mM–400 mM NaCl was used to elute bound proteins. BFD eluted from the column at 280–300 mM NaCl. For further purification, hydrophobic interaction chromatography (HIC) was employed. Prior to loading the protein on the column, NaCl was removed and the sample volume reduced by ultrafiltration. Ammonium sulfate was added to a final concentration of 0.25 M before the protein was loaded onto a Phenylsepharose high sub. column (Pharmacia) that had been equilibrated with 50 mM potassium phosphate buffer containing 0.25 M ammonium sulfate. Elution of BFD was accomplished using a linear gradient of 0.25–0.0 M ammonium sulfate. BFD eluted at the end of the gradient (0.05–0 M) and the active fractions were pooled, desalted by ultrafiltration, and lyophilized. C. Construction, Expression, and Purification of the BAL Variants The C-terminal 6x-his-tagged BAL expression vector, pKKBAL-his, was available from a previous study (22). The vectors pKKBALA28S, pKKB-


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ALQ113A, and pKKBALQ113H were generated using the methodology described earlier for the BFD mutants. The forward primers used for mutagenesis were: A28S: 5V-GTTCGGCCTGCACGGatCcCATATCGATACG-3V Q113H: 5V- GAAACCAAtACGTTGCAcGCGGGGATTGATCAGG-3V Q113A: 5V-GATGAAACCAACACGcTagcGGCGGGGATTGATCAGG-3V For A28S this adds a BamH1 restriction site, which allows for ready screening of transformants. For Q113A an Nhe1 site was added, while for Q113H an AflIII site is lost. The expression and purification of BAL-his variants was accomplished using essentially the same procedure as for the PDC-his variants described earlier. The storage buffer for the BAL variants was 50 mM potassium phosphate pH 7.0, containing 1 mM MgSO4, 0.5 mM TDP, and 10% glycerol. All protein concentrations were determined by using the Bradford method (29), with BSA as the standard. D. Assay of Decarboxylase Activity The decarboxylation of pyruvate was measured using a coupled enzymatic assay, as described elsewhere (30). The continuous photometric assay was performed at 30jC in 50 mM Mes/KOH, pH 6.5, 0.1 mM TDP, 2 mM MgSO4, 30 mM 2-keto acid, 0.035 mM NADH, and 0.05 U horse liver alcohol dehydrogenase (HLADH). One unit is defined as the amount of PDC that catalyzes the decarboxylation of 1 Amol keto acid per minute at pH 6.5 at 30jC. A continuous coupled photometric assay has been described for BFD (31). In this study the assay was performed at 30jC in 50 mM potassium phosphate buffer, pH 6.0, 0.1 mM TDP, 2.5 mM MgSO4, 30 mM 2-keto acid, 0.035 mM NADH and 0.05 U HLADH. One unit is defined as the amount of BFD that catalyzes the decarboxylation of 1 Amol keto acid per minute at pH 6.0 at 30jC. E. Assay of Benzaldehyde Lyase Benzaldehyde lyase was also assayed using the same assay as employed for BFD. The assay mixture contains 50 mM Tris pH 8.0, 1 mM MgSO4, 0.5 mM TDP, 15% PEG 400, 0.035 mM NADH, and 0.10 units HLADH. The substrate, benzoin, was prepared in 50 mM Tris pH 8.0 containing 15% PEG. One unit is defined as the amount of BAL that catalyzes the cleavage of 1 Amol benzoin per minute at pH 8.0 at 30jC.

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III. RESULTS AND DISCUSSION Amino acid sequences of BAL (AY007242), BFD (1BFD), ZmPDC (1ZPD), and ScPDC (1PVD) were obtained from either GenBank (BAL) or the PDB. Alignments of the sequences showed that the proteins shared about 25% sequence identity and 35% sequence similarity, with, not surprisingly, the two PDCs being more similar to each other. The two residues previously suggested to play a role in the binding of pyruvate to ZmPDC, Ile472 and Ile476 (25), had counterparts in Ile476/480 in ScPDC, Ala460 and Phe464 in BFD, and Ala480 and Phe484 in BAL. The structures of BFD (11), ZmPDC (12), and ScPDC (32) were also superimposed, confirming the suggestion that Ala460/ Phe460 were indeed the BFD analogs of Ile472/476. Models of BAL were prepared using each of 1BFD, 1PDC, and 1ZPD as a template. 1ZPD was possibly the best template, but the structures of the active site look essentially identical for all models. Figure 2 shows portions of a combined structural/sequence alignment of BAL with the three decarboxylases; a model of the active site of BAL is shown in Figure 3. The overall alignments show that 22 residues are conserved among all these enzymes, including the active-site glutamate and the metal-binding motif. Of particular interest is the observation that BAL does not contain the histidine residues common to the decarboxylases. Instead, BAL contains a glutamine residue, and its NE2 superimposes on the NE2 of His70 (1BFD) and His114 (1ZPD). In this respect BAL is more similar to pyruvate oxidase from Lactobacillus plantarum, which has a glutamine positioned similarly in its active site (33). Both BAL and BFD bind aromatic substrates, and it is no surprise that the residues thought to bind the aromatic portion of the substrate were identical, i.e., Ala and Phe, whereas the two PDCs had isoleucine residues at the corresponding positions. One potentially significant change in BAL was the replacement of Ser26 (BFD) and Asp27 (ZmPDC) by an alanine residue (Ala28). In addition to substrate binding, both Ser26 (15) and Asp27 (18,34) have been implicated in the catalytic mechanism of the decarboxylases. It is conceivable that this substitution facilitates the binding of the benzoin substrate while, at the same time, reducing any propensity BAL might have for decarboxylase activity. Based on these alignments (Figs. 2 and 4) and the results of Pohl et al. (25), it was decided that the first approach toward interconverting the substrate specificity of BFD and PDC would be via the replacement of ZmPDC Ile472 and Ile476 with Ala and Phe, respectively. Concomitantly, BFD Ala460 and Phe464 would be replaced by Ile. ZmPDC was chosen as the PDC for mutagenesis because its substrate specificity was much narrower than that of ScPDC and it is unable to decarboxylate benzoylformate (24). These substitutions were carried out in a stepwise manner; the purification


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Figure 2 Portions of the combined sequence and structural alignment of BFD (PDB 1BFD), ZmPDC (PDB 1ZPD), ScPDC (PDB 1PVD), and BAL. Residues fully conserved across all enzymes are in boldface while the essential glutamic acid residue and the metal binding residues are in boldface and underlined. The histidine residues implicated in catalysis by BFD and PDC, and the active site glutamine of BAL, are arrowed (;) while residues involved in substrate binding are also indicated (d).

and specific activities of wild-type (WT) and variant PDC and BFD are shown in Tables 1 and 2, respectively. It was possible to readily purify all variants of both enzymes by the same method used for the WT, suggesting that the mutagenesis did not result in gross conformational changes. Clearly there was some loss of specific activity, with the activity of the double mutant dropping to about 2–3% of that of the WT. Table 3 shows the substrate specificity of WT and mutant PDC and BFD. In line with earlier observations (24), PDC was found unable to decarboxylate benzoylformate. The I472A mutation resulted in a twofold decrease in pyruvate decarboxylase activity but also in considerably enhanced

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Figure 3 Model of the active site of BAL based on the sequence/structure alignments shown in Figure 2. (See color insert.)

Figure 4 Overlay of active-site residues of BAL with those of (A) BFD and (B) ZmPDC. (See color insert.)


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Table 1 Purification of WT ZmPDC-His and ZmPDC-His Mutantsa Enzyme WT-PDC PDCI472A PDCI476F PDCI472A/I476F a

Crude extract (U/mg)

Ni-NTA, gel filtration (U/mg)

30 10 2.7 1.3

120–140 40–50 20–25 2–3

Specific activities were obtained using the standard PDC assay described in Section II.

activity toward longer-chain aliphatic substrates. More importantly, it was able to decarboxylate benzoylformate at about 3% of the rate at which pyruvate was decarboxylated. In some respects the I476F mutation had the most deleterious impact on the decarboxylation rate of all substrates, because all rates decreased and no selectivity for longer-chain length was observed. However, this variant did also have some BFD activity, albeit at a low level. The I472A/I476F double mutant showed the greatest decrease in pyruvate decarboxylase activity and the greatest selectivity for long-chain aliphatic substrates. For example, WT ZmPDC is effectively unable to decarboxylate a C6 keto acid, a task the I472A/I476F double mutant is able to accomplish 20 times more rapidly than it is able to decarboxylate pyruvate. That given, in the context of this work the most notable observation was that, although the overall rate was low, the double mutant was able to decarboxylate benzoylformate at about 25% of the pyruvate decarboxylation rate. The BFD variants also showed significant changes in activity across the substrate spectrum. Wild-type BFD possesses very little PDC activity (less than 0.01% of its BFD activity), and none of the variants constructed here provided a major improvement in absolute rate. The best was A460I, which was able to decarboxylate pyruvate eight times faster than was WT BFD.

Table 2 Purification of WT BFD and BFD Variantsa

BFD BFDA460I BFDF464I BFDA460I/F464I a

Crude extract

Anion exchange

Hydrophobic interaction

73 1.1 1.9 0.4

270 12 28 3.6

360 17 33 5.6

Specific activities, reported in units per milligram, were obtained using the standard BFD assay described in Section II.

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Table 3 Specific Activity of ZmPDC and BFD Variants with Aliphatic and Aromatic 2-Keto Acidsa

a

Specific activities are reported in units per milligram. Data obtained using the standard PDC assay. c Data obtained using the standard BFD assay. b

Nonetheless, even that rate was only 0.25% of its rate of benzoylformate decarboxylation. As with the PDC variants, there was an increase in capacity to decarboxylate the longer chain aliphatic keto acids, particularly with the A460I mutant. Indeed, BFD A460I is able to decarboxylate the C6 keto acid at a rate not dissimilar to that shown by PDC I472A. Overall, although the absolute rates are faster for the pyruvate decarboxylases, the mutations in both enzymes show broadly the same trends, with the major difference between the two enzymes being that the A460I mutation in BFD resulted in a greater decrease in WT activity (23-fold) than the F464I mutation (ninefold). Conversely, in PDC the decrease in activity was more significant for the I476F variant (sixfold) than the I472A variant, which showed about a twofold decrease. In toto these results suggest that we have gone part of the way toward interconverting the substrate specificity of the two enzymes. While we have been successful at increasing the level of BFD activity in ZmPDC, and vice versa, the absolute rates are not particularly good. However, using either enzyme as the template, we have been able to construct a C6 keto acid decarboxylase with reasonable activity. This suggests that the PDC active site needs further manipulation to optimize binding of aromatic substrates and that the BFD active site is still too large to optimally bind small aliphatic


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substrates. Moreover, it also suggests that considerably more than the point mutations described here will be required to achieve this. In addition to its decarboxylase action, BFD is able to carry out the stereospecific synthesis of R-benzoin (35,36). Given that BAL catalyzes the breakdown of R-benzoin and shares at least two substrate recognition residues with BFD (Fig. 4A), it seemed appropriate to see whether these two enzymes could be also be interconverted. Two potentially significant differences in the active sites of these enzymes are the substitution of Ser26 and His70 in BFD by Ala28 and Glu113, respectively, in BAL (Fig. 4A). Accordingly, BFD S26A and H70Q as well as BAL A28S and Q113H were prepared. Further, it has been shown that the replacement of His70 by alanine leads to a decrease in the activity of BFD of more than three orders of magnitude (15,37). It was of interest to see whether the same substitution in BAL would have a comparable effect, so the BAL Q113A variant was also prepared. All BAL-his variants were prepared using basically the same purification scheme as for the PDC variants, with results similar to those in Table 1 (data not shown). The Km and kcat values for WT BAL-his were 60 lM and 88 s1, respectively. The kinetic parameters obtained following the mutagenesis of BAL and the Km and kcat values obtained for the corresponding BFD variants are all shown in Table 4. The BAL A28S mutation provided a 10-fold increase in the Km value for R-benzoin but only a sixfold decrease in lyase activity. This can be compared to a 23-fold increase in the Km value for benzoylformate and greater than a 50-fold decrease in kcat for the BFD S26A variant. Ser26 has been implicated in several stages of the catalytic cycle of BFD (15,37), and it appears to be relatively more important for BFD than Ala28 is for BAL. It is possible that the changes in lyase activity brought about by the A28S mutation may simply be related to the greater bulk of the serine residue, making it slightly more difficult to bind the substrate and position it for catalysis. The H70A substitution in BFD led to a decrease in kcat of more than three orders of magnitude as well as an increase in Km for benzoylformate (15,37). This suggests that His70 is crucial for BFD activity. The comparable mutation in BAL, Q113A, results in little change in Km for R-benzoin and a decrease in kcat of 160-fold. Clearly, while Gln113 is important for BAL, it is not essential for lyase activity. Not unexpectedly, replacement of BFD His70 by glutamine and BAL Gln113 by histidine resulted in decreased activity for both enzymes. However, the decrease in kcat for BAL Q113H (120-fold) was similar to the decrease observed for the Q113A variant. By contrast, the H70Q mutation in BFD led to a 250-fold decrease in the value of kcat, a substantially smaller decrease than that observed for the H70A variant (3500fold) but with essentially no change in the value of Km. Sequence and structural alignments suggest that BAL, pyruvate oxidase (33), and acetolactate

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Table 4 Lyase and Decarboxylase Activity of BAL and BFD Variantsa

BAL

BFD

Km benzoin (mM)

kcat BAL (s1)

Km benzoylformate (mM)

kcat BFD (s1)

WT S26A H70A H70Q

0.06 0.73 0.04 0.03 — n.a.c n.a.c —

87.6 14.3 0.54 0.74 n.d.b n.a.c n.a.c n.d.b

— 17.4 F 1.0 n.a.c — 0.37d 8.6d 1.9d 0.2

n.d.b 1.7 F 0.1 n.a.c n.d.b 241d 4.5d 0.07d 0.94

WT A28S Q113A Q113H

a

Michaelis–Menten parameters for lyase activity were obtained using the BAL assay (pH 8.0) with varying concentrations of benzoin. The parameters for benzoylformate decarboxylase activity were obtained using the BFD assay (pH 6.0) with varying concentrations of benzoylformate. b n.d.; no activity detected using the standard spectrophotometric coupled assay. It has been reported that BAL does have a detectable level of decarboxylase activity when HPLC is used to measure the production of benzaldehyde (Ref. 41). c n.a.; not assayed. d From Polovnikova et al. (Refs. 15,37).

synthase (38) all have a glutamine residue located near the 4V-imino group of TDP. In BFD and both ZmPDC and ScPDC, a histidine residue replaces the glutamine. Perhaps the most interesting case is that of transketolase, in which the bacterial and yeast enzymes have a histidine (His481) adjacent to the 4Vimino group (39). In the mammalian enzyme, the histidine is replaced by a glutamine (40). It is not yet clear why the individual enzymes have a preference for histidine or glutamine, but certainly the question warrants further investigation. Although the BAL A28S mutation did not produce significant changes in benzaldehyde lyase activity, it did produce the most surprising result. As suggested earlier, both BAL and BFD carry out identical carboligation reactions and share similar residues for substrate recognition. However, as shown in Table 4, BFD cannot cleave R-benzoin, nor is any benzoylformate decarboxylase activity seen with BAL. Replacement of His70 in BFD by glutamine or Gln113 in BAL by histidine has no effect on the reaction profile for either enzyme. However, the A28S variant of BAL does possess benzoylformate decarboxylase activity. Admittedly, the Km for benzoylformate is relatively high (17 mM), but the activity is readily measurable by the spectrophotometric coupled assay, with a kcat value of 1.7 s1. There has been only


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one previous report of BAL possessing benzoylformate decarboxylase activity. A sensitive HPLC assay was necessary to measure the benzaldehyde produced, and it provided a kcat value of less than 0.02 s1 (41). Thus the BAL A28S mutation provides at least a 100-fold increase in BFD activity and lends further credence to suggestions that Ser26 in BFD plays a critical role in catalysis by BFD (15,37).

IV. SUMMARY Benzoylformate decarboxylase, pyruvate decarboxylase, and benzaldehyde lyase are all thiamine diphosphate–dependent enzymes that either catalyze similar reactions (BFD and PDC) or catalyze reactions that result in similar products (BFD and BAL). All these enzymes are also able to carry out comparable carboligation reactions, but with an altered spectrum of substrates. Further, the X-ray structures show that the active sites of BFD and PDC are also very alike. Given these similar features, it is of interest to identify those residues that contribute to the differences between the enzymes. Here we use a combination of sequence and structural alignments to prepare a homology model for BAL and to identify some of the residues potentially involved in substrate binding and specificity for each of these enzymes. Sitedirected mutagenesis is then used to construct variants of each enzyme that possess some of the residues of their counterparts. The results show that it is possible to make relatively simple changes that will alter the substrate specificity of each of these enzymes. We have been able to observe pyruvate decarboxylase activity with BFD variants and benzoylformate decarboxylase activity with PDC variants. Perhaps the most intriguing result is that we have been able, with a single point mutation, to convert benzaldehyde lyase into a benzoylformate decarboxylase.

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involved in catalysis by Zymomonas mobilis pyruvate decarboxylase. Biochem J 339:255–260, 1999. B Gonza´lez, R Vicunã. Benzaldehyde lyase, a novel thiamin PPi-requiring enzyme, from Pseudomonas fluorescens biovar I. J Bacteriol 171:2401–2405, 1989. P Hinrichsen, I Go´mez, R Vicunã. Cloning and sequencing of the gene encoding benzaldehyde lyase from Pseudomonas fluorescens biovar I. Gene 144:137–138, 1994. AS Demir, M Pohl, E Janzen, M Muller. Enantioselective synthesis of hydroxy ketones through cleavage and formation of acyloin linkage. Enzymatic kinetic resolution via C–C bond cleavage. J Chem Soc Perkin Trans 1:633–635, 2001. H Iding, P Siegert, K Mesch, M Pohl. Application of a-keto acid decarboxylases in biotransformations. Biochim Biophys Acta 1385:307–322, 1998. GA Sprenger, M Pohl. Synthetic potential of thiamin diphosphate–dependent enzymes. J Mol Catalysis B: Enzymatic 6:145–159, 1999. M Pohl, P Siegert, K Mesch, H Bruhn, J Grotzinger. Active-site mutants of pyruvate decarboxylase from Zymomonas mobilis—a site-directed mutagenesis study of L112, I472, I476, E473, and N482. Eur J Biochem 257:538–546, 1998. SN Ho, HD Hunt, RM Horton, JK Pullen, LR Pease. Site-directed mutagenesis by overlap extension using the polymerase chain reaction. Gene 77:51–59, 1989. EA Sergienko, J Wang, L Polovnikova, MS Hasson, MJ McLeish, GL Kenyon, F Jordan. Spectroscopic detection of transient thiamin diphosphate–bound intermediates on benzoylformate decarboxylase. Biochemistry 39:13862–13869, 2000. AY Tsou, SC Ransom, JA Gerlt, DD Buechter, PC Babbitt, GL Kenyon. Mandelate pathway of Pseudomonas putida: sequence relationship involving mandelate racemase, (S)-mandelate dehydrogenase, and benzoylformate decarboxylase and expression of benzoylformate decarboxylase in Escherichia coli. Biochemistry 29:9856–9862, 1990. MM Bradford. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72:248–254, 1976. M Pohl, J Grotzinger, A Wollmer, MR Kula. Reversible dissociation and unfolding of pyruvate decarboxylase from Zymomonas mobilis. Biotechnol Appl Biochem 22:95–105, 1994. PM Weiss, GA Garcia, GL Kenyon, WW Cleland, PF Cook. Kinetics and mechanism of benzoylformate decarboxylase using 13C and solvent deuterium isotope effects on benzoylformate and benzoylformate analogues. Biochemistry 27:2197–2205, 1988. F Dyda, W Furey, S Swaminathan, M Sax, B Farrenkopf, F Jordan. Preliminary crystallographic data for the thiamin diphosphate–dependent enzyme pyruvate decarboxylase from brewer’s yeast. J Biol Chem 265:17413–17415, 1990. YA Muller, G Schumacher, R Rudolph, GE Schulz. The refined structures of a stabilized mutant and of wild-type pyruvate oxidase from Lactobacillus plantarum. J Mol Biol 237:315–335, 1994.

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34. EA Sergienko, F Jordan. Catalytic acid–base groups in yeast pyruvate decarboxylase. 2. Insights into the specific roles of D28 and E477 from the rates and stereospecificity of formation of carboligase side products. Biochemistry 40: 7369–7381, 2001. 35. AS Demir, T Du¨nnwald, H Iding, M Pohl, M Mu¨ller. Asymmetric benzoin reaction catalyzed by benzoylformate decarboxylase. Tetrahedron: Asymmetry 10:4769–4774, 1999. 36. H Iding, T Du¨nnwald, L Greiner, A Liese, M Mu¨ller, P Siegert, J Gro¨tzinger, AS Demir, M Pohl. Benzoylformate Decarboxylase from Pseudomonas putida as stable catalyst for the synthesis of chiral 2-hydroxy ketones. Chem Eur J 6:1483–1495, 2000. 37. L.S. Polovnikova, Investigation of thiamin diphosphate–dependent catalysis by benzoylformate decarboxylase. PhD dissertation, Purdue University, West Lafayette, IN, 2000. 38. D Chipman, Z Barak, JV Schloss. Biosynthesis of 2-aceto-2-hydroxy acids: acetolactate synthases and acetohydroxyacid synthases. Biochim Biophys Acta 1385:401–419, 1998. 39. S Ko¨nig, A Schellenberger, H Neef, G Schneider. Specificity of coenzyme binding in thiamin diphosphate–dependent enzymes. crystal structures of yeast transketolase in complex with analogs of thiamin diphosphate. J Biol Chem 269: 10879–10882, 1994. 40. G Schenk, R Layfield, JM Candy, RG Duggleby, PF Nixon. Molecular evolutionary analysis of the thiamine diphosphate–dependent enzyme, transketolase. J Mol Evol 44:552–572, 1997. 41. E. Janzen. Die Benzaldehydlyase von Pseudomonas fluoreszenz. PhD dissertation, Heinrich Heine University, Du¨sseldorf, Germany, 2002.

17 Benzoylformate Decarboxylase: Intermediates, Transition States, and Diversions Ronald Kluger, Qingyan Hu, and Ian F. Moore* University of Toronto, Toronto, Ontario, Canada

I. INTRODUCTION The properties of reactions involving intermediates derived from thiamine have provided quantitative comparisons with the combination of thiamine diphosphate and a protein in enzymic catalyzed reactions (1,2). Where both thiamine and TDP enzymes catalyze similar processes, we can compare the rates of individual steps, specificity of the reactions, and competitive processes (2–4). This chapter deals with the effects of a surprising alternative to a wellknown process in thiamine catalysis and an appreciation of the hidden challenges that the enzyme overcomes. II. INTERMEDIATES IN BENZOYLFORMATE DECARBOXYLASE Benzoylformate decarboxylase (BFD), an enzyme in the mandelate pathway, converts benzoylformate to carbon dioxide and benzaldehyde (5–8). It utilizes

*Current affiliation: McMaster University, Hamilton, Ontario, Canada

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Scheme 1

thiamine diphosphate (TDP) as a cofactor, presumably forming the covalent addition intermediate a-mandelylthiamin-DP (MTDP). Loss of carbon dioxide from MTDP produces the conjugate base of (hydroxybenzyl)thiamine-DP (HBzTDP). This intermediate must be protonated at carbon prior to the release of benzaldehyde, regenerating the enzyme and cofactors (Scheme 1). Loss of the proton from the hydroxyl group of HBzTDP leads to elimination of the benzaldehyde and regeneration of the TDP.

III. HYDROXYBENZYLTHIAMIN (HBzT): ANALOG OF THE BFD INTERMEDIATE The central intermediates in catalysis by BFD are the addition products, MTDP and HBzTDP. In order to learn the functions of the protein in BFD (as distinct from TDP itself), it is important to observe the nonenzymic reactivity of the corresponding covalent intermediates (3). Since the diphosphate of the coenzyme is known to be remote from the reaction site (7), where it serves an enzyme-binding role, reactivity studies are more readily carried out with derivatives where thiamine replaces TDP (2,4,9). The combination of benzaldehyde and thiamine under basic conditions produces HBzT. The reaction that has been studied since the condensation of benzaldehyde with itself to produce benzoin is catalyzed by thiamine in basic

BFD: Intermediates, Transition States, and Diversions

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Scheme 2

solutions, proceeding via HBzT as an intermediate (10–14). Alternatively, HBzT should result from decarboxylation of a-mandelylthiamin, in analogy to the reaction that produces HBzTDP on BFD, a route that had not been achieved in a nonenzymic reaction (Scheme 2). The reactions and generation of HBzT are not as simple as these schemes suggest and revelations of the role of the protein in BFD arise from the comparison. IV. HBzT UNDERGOES C2a a PROTON EXCHANGE Pioneering NMR studies by Hank Sable and his coworkers showed that HBzT undergoes proton-deuteron exchange in deuterium oxide at C2a (Scheme 3) (15,16). The probable intermediate is the conjugate base of HBzT at C2a. Sable called this intermediate ‘‘the second carbanion in thiamine catalysis,’’ a species that has an enamine resonance contributor.

V. HBzT DOES NOT PRODUCE BENZALDEHYDE IN NEUTRAL SOLUTION The conversion of HBzT to benzaldehyde and thiamine is the reverse of its established synthesis. The synthesis of HBzT is accomplished under basic

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Scheme 3

conditions, necessary to provide a significant concentration of the C2 conjugate base of thiamine (the thiamine ylide). In basic solution in the presence of excess benzaldehyde, HBzT transfers the C2a proton. The resulting carbanion adds to the carbonyl carbon of benzaldehyde to give the conjugate of benzoin. In the absence of added benzaldehyde, the carbanion undergoes an alternative reaction, an irreversible fragmentation that destroys thiamine (17,18). The reaction gives dimethyl amino pyrimidine (DMAP) and a phenyl thiazole ketone (PTK). The alternative reaction is the reverse of the synthesis of HBzT, loss of benzaldehyde after ionization of the hydroxyl group at C2a (Scheme 4). However, at pH 7 the rate of the fragmentation process (followed by the absorbance of PTK at 328 nm) is 1000 times the rate

Scheme 4


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of release of benzaldehyde (extrapolated from measurements at higher pH) (17,19). VI. N-ALKYLATION OF HBzT LOCALIZES CHARGE We studied the pH dependence of the fragmentation reaction and found that the rate is proportional to the extent of protonation of its pyrimidine group. By alkylation of N1 of the pyrimidine, we created a positive charge that is not subject to ionization. N1-Methylated HBzT undergoes fragmentation to dimethyl N-methyl amino pyrimidine (DMP) and PTK, even in alkaline solutions (17). We also observe that the overall fragmentation process is buffer catalyzed and that the corresponding N-benzylated derivative also reacts in the same way (19) (Scheme 5). VII. NMR ANALYSIS OF FRAGMENTATION AND EXCHANGE IN N1-METHYL HBzT The fragmentation of N1-methyl HBzT into DMP and PTK can be followed by proton NMR, since chemical shifts change in the fragmentation. The exchange of the C2a proton for deuterium in deuterium oxide is followed at the same time as the intensity of that peak decreases as the proton is replaced, as shown in scheme 6. We observe that HBzT fragments more slowly than it undergoes H–D exchange at C2a exchange (17). This is consistent with Sable’s observation of the exchange reaction (15,16). However, as the reaction proceeds, the rate of fragmentation decreases. This is the result of the substrate’s becoming deuterated, acquiring a deuteron faster than it undergoes fragmentation (kBH [BH] >kf). Since proton removal is partially rate limiting for fragmentation, there should be a primary isotope effect on the observed reaction rate.

Scheme 5

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Scheme 6

VIII. FRAGMENTATION OF HBzT IS CATALYZED BY HYDROXIDE AND BY ANIONIC BRØNSTED BASES We studied the effect of buffers on the rate of the fragmentation reaction of the N1-benzyl derivative of HBzT by following the appearance of the PTK (19). The rate plot reveals a first-order dependence on hydroxide. Examination of buffer catalysis reveals a route that involves general base catalysis (by anionic bases only; Brønsted h = 0.5). This is consistent with the fact that proton transfer from carbon is completely rate determining, in disagreement with the NMR results that suggest that the fragmentation is at least partially rate determining (using the N1-methyl derivative—the compounds are similar in reactivity) (17). However, the UV method we used for obtaining these data were done with much more dilute buffers than were used the NMR studies (NMR requires higher substrate concentrations, necessitating higher buffer concentrations). Therefore, we examined the effects of changes in the concentration of buffer on the rate of the reaction in order to determine the source of the inconsistency (19).

IX. THE EFFECT OF THE BUFFER IS SUBJECT TO SATURATION: CHANGE IN RATE-DETERMINING STEP The discrepancy between the sets of conditions can be reconciled by the information in a plot of observed rate at pH 6.1 as a function of the concentration of the phosphate buffer. At high buffer concentrations, the rate is


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Figure 1 Dependence of the rate of fragmentation on phosphate buffer concentration at pH 6.1 (or the corresponding pD). The upper curve is for the reaction in deuterated water.

Scheme 7

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independent of buffer concentration; at low buffer concentrations, the rate increases with increasing buffer concentration. The rate law is kobs ¼ ðkB ½B þ kOH ½OH Þkf = kBH ½BH þ kw þ kf The overall rate is faster in deuterium oxide, giving an inverse solvent isotope effect (see Fig. 1) with k(DOD)/k(HOH) = 3.0 at saturation. This result is typical of a reaction that proceeds by an E1CB mechanism: The carbanion intermediate is more slowly quenched by deuterium oxide and by deuterated acids (kBD) than by water and protic acids (20). At low buffer concentrations, proton transfer from C2a is rate-limiting; increasing the concentration of buffer enhances the observed rate of fragmentation. At high buffer concentrations, fragmentation is rate limiting, and further increases in buffer concentration have no effect (Scheme 7). The nonlinearity of the dependence enabled us to calculate the rate constant for the fragmentation step based on an estimate of the pKa for loss of a proton from the C2a position of HBzT: kf is about 105 s1 at 40jC. Thus, we know that the fragmentation step itself is very fast and comparable to the rate of protonation of the intermediate. The process can be represented by electron movement that transfers the proton from the hydroxyl group to the methylene bridge (Scheme 8). However, this is requires a transition state that would be very high in energy since it involves unusual orbital occupations. An alternative, which is more easily seen from the carbanion resonance structure, is an internal hydride shift (Scheme 9). We have studied the mechanism in more detail, and the results will be published. The competition provided by the very low barrier to this process is a significant problem that the enzyme has overcome.

Scheme 8


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X. a -(MANDELYL)THIAMINE—THE INTERMEDIATE IN THIAMINE-CATALYZED DECARBOXYLATION OF BENZOYLFORMATE At this point we know of the complex reaction patterns of HBzT and its conjugate base. In order to calibrate the reaction sequence in BFD, we also need to evaluate the reaction patterns of the precursor in the decarboxylation process, a-mandelyl-thiamine (MT), which should be accessible through synthesis. Our first attempts at preparing this material were unsuccessful. We had reasoned that the preparation of a-lactyl-thiamine, starting from ethyl pyruvate and the conjugate base of thiamine, would be the obvious model. We would combine the conjugate base of thiamine with ethyl benzoylformate to produce the ethyl ester of MT and then hydrolyze the ester. However, our attemps to accomplish the condensation by this process were unsuccessful. Having observed that magnesium chloride polarizes carbonyl groups in another study, we added magnesium chloride to catalyze the addition, with success. The ester hydrolyzes in concentrated hydrochloric acid (Scheme 10).

Scheme 10

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In strong acid, decarboxylation is suppressed, since the COOH must ionize for reaction to occur. The rest of the molecule is also stable in acid.

XI. KINETICS OF DECARBOXYLATION OF a -(MANDELYL)THIAMINE We measured the rate of decarboxylation of MT (Scheme 11) by following changes in its UV spectrum. The pH dependence of the rate data fits for carboxylic acid group having a pKa of about 0.2, indicating that MT is unusually acidic for a carboxylic acid. Since both the internal pyrimidine and thiazolium are cationic in acid, the dissociation of the carboxyl to form an anion is significantly stabilized. The pH-rate data also fit a pKa of about 3.5, which is likely to result from ionization of the protonated pyrimidine. The value is close to that found by titration of thiamine. The first-order rate constant for decarboxylation of MT at 25jC, under conditions where the pyrimidine is protonated and the carboxyl is not protonated, is 5.8 104 s1, corresponding to a half-life of about 20 minutes. At higher pH (5–8), where both groups are not protonated, the observed first-order rate constant is 3.7 104 s1, which is a half-life of about 31 minutes. For comparison the catalytic rate constant of BFD is 81 s1 (20a), indicating that the enzyme accelerates the rate of decarboxylation of the intermediate by a factor of at least 105. While the nonenzymic rate for MT is slow, the decarboxylation of benzoylformate must be much slower: It is so slow that cannot be detected. This shows that formation of MT itself enhances the rate of decarboxylation to a very large degree and that the enzyme adds a relative small acceleration, lowering the activation barrier by about 4 kcal. Since the decarboxylation process is unimolecular, the enzyme cannot impart an entropic advantage. Since the reaction cannot be subject to acid/base catalysis (carbon dioxide

Scheme 11


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must leave prior to protonation at the site), the enzyme is further limited in devices that could be used to achieve this acceleration. Lienhard showed that in general this class of reaction is accelerated in a medium of lower polarity than water, because the zwitterionic reactant passes through a transition state with much less charge separation (21,22), and he used this information to design an inhibitor of pyruvate dehydrogenase (23). However, that leaves the enzyme with an energy problem: If the covalent intermediate, MTDP, is more stable in water and benzoylformate is initially dissolved in water, then there is an energetic price for the desolvation that reduces any kinetic acceleration. A similar problem exists with the addition of pyruvate to TDP enzymes (9). The enzyme must have the capability of coupling energy released in forming the intermediate and/or binding the substrate to accomplish the desolvation.

XII. FRAGMENTATION PRODUCTS FROM DECARBOXYLATION OF MT The loss of carbon dioxide from MT should produce the conjugate base of HBzT, the first species involved in the fragmentation process (Scheme 12). In that case, the intermediate is generated by the transfer of a proton to an acceptor. Here, the loss of carbon dioxide is unimolecular. Based on the fragmentation patterns of HBzT (17,19), we predicted that the intermediate formed by loss of carbon dioxide should fragment to PTK and DMAP, while Brønsted acids should divert it to HBzT (Scheme 13). Therefore, we examined the absorbance spectrum product for absorbance centered at 328 nm, which would arise from the fragmentation product, PTK (Scheme 13). Spectra recorded over several hours reveal that the rate of appearance of PTK coincides with the rate of decarboxylation, consistent with the expected fragmentation (Fig. 2).

Scheme 12

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Scheme 13

XIII. BUFFER CHANGES YIELD, NOT RATE CONSTANT As already stated, the carbanion resulting from loss of carbon dioxide from MT is the same species that results from the loss of a proton from HBzT. The rate constant for the fragmentation of the conjugate base of HBzT at 40jC is 105 s1 (24), which will give a rate that is comparable to that for protonation of a model related to this intermediate. Therefore, we expect that at low buffer concentrations, the yield of PTK relative to HBzT from

Figure 2 Spectra recorded for solutions of MT at 25jC, pH 7.0.


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MT will be highest. We observe this to be the case (Fig. 3). The observed rate coefficient for decarboxylation of MT is not affected by buffer concentration, but the relative amount of PTK is affected. The buffer decreases the yield of PTK. Protonation of the intermediate by water, which is a very weak acid, is very slow and does not compete with the proton transfer by stronger Brønsted acids. The rate plot at two buffer concentrations makes this apparent. Moreover, the effect of buffer on product distribution can be made into a quantitative result. We know that HBzT is produced by protonation of the intermediate. However, HBzT can undergo the reverse reaction and regenerate the intermediate, which can fragment or reprotonate. There is an initial burst of PTK formation and then a slow transformation of HBzT into PTK until all the HBzT is consumed. At the end, MT is completely converted to PTK. Therefore, we measured the absorbance of the solution of MT after several hours to obtain the absorbance of the initially formed PTK. Then we heated the solution until no further reaction occurred. The amount of PTK

Figure 3 Change in absorbance of solutions of MT at high buffer concentration (lower curve) and low buffer concentrations (upper curve). The observed rate constants are the same, but the yields of the absorbing species (PTK) are inversely proportional to buffer concentration.

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generated in this second process is equal to the amount HBzT that originally formed. The total amount of PTK (and DMAP) produced is equal to the original amount of MT, so the ratio of the final absorbance at 328 nm compared to the absorbance at the end of the first reaction gives the fraction of HBzT formed initially (in the total amount of PTK and HBzT). Using varying buffer concentrations, we fit the yield equation to calculate the ratio of rates of formation of PTK and HBzT from the intermediate (Fig. 4). The inherent tendency of the intermediate to fragment is overcome by the Brønsted acid component of the buffer. The loss of carbon dioxide from MT requires rehybridization of the departing carboxyl from sp2 to sp as well as lengthening of the C–C bond. Once CO2 is separated, fragmentation competes with protonation. The rehybridization process is likely to be concerted with the bond-breaking step. Guthrie has presented a detailed analysis for the case of the release of carbon dioxide from acetoacetic acid (25). This requires that protonation occur only after the carbon dioxide molecule is fully formed (Scheme 14). In that case, fragmentation can begin prior to the second-order process that would be required for protonation. Furthermore, the inherent rate for protonation of this type of species by water is very slow, although the species is more basic than hydroxide, so fragmentation has little inherent competition. Thus, since fragmentation in solution is unimolecular and rapid, the enzyme must avoid this pathway to prevent destruction of TDP. However, evidence from Jordan’s group indicates that protonation is not accelerated (8). Jordan and coworkers observed that when p-nitrobenzoylformate reacts with BFD, a

Figure 4 Yield of HBzT as a function of buffer concentration where the alternative product is PTK.


Scheme 14

Scheme 15

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stable absorbance develops at 410 nm, which they assign to the carbanionenamine intermediate (Scheme 15). This species should fragment with a rate constant that would be much too large to allow the intermediate to be observed, yet the absorbance persists. Alternatively, if fragmentation were normally avoided on the enzyme by rapid protonation of the intermediate, then the absorbance would be quenched even more rapidly than fragmentation could occur. Since Jordan observes that the enamine’s aborbance persists (the rate constant for its disappearance is 4.4 s1 while that for fragmentation at this temperature would be 1000 times greater), proton transfer to nitro-HBzT must not be facilitated (Scheme 16). If the absorbance is indeed from the intermediate, then the enzyme must prevent the inherent tendency to fragment without rapid protonation. At this point we can speculate as to how the enzyme can block the low barrier to fragmentation without facilitating protonation. It appears that the enzyme has met the challenge not simply by favoring the protonation route but by avoiding the fragmentation route completely. Any bimolecular process would not overcome the unimolecular competition completely. Therefore, we expect that the enzyme provides specificity for the protonation route by effectively raising the barrier to the fragmentation route. This is reminiscent of the specificity issues in competing routes from a common intermediate in enzymes that generate covalent intermediates in PLP-dependent enzymes. Dunathan proposed that stereoelectronic barriers can be created by avoiding conformations that would lead to alternative routes (26). Basically, the pathways that are blocked have orbitals that cannot align to delocalize as the intermediate is held in a specific conformation by the protein. In the case of BFD we might expect to find a similar utilization of stereoelectronic control. However, we do not know the mechanism of the fragmentation step, and therefore we do not know which overlaps would be avoided. With this in mind it will be interesting to learn the details of the alignment of the substrate in the active site and also to learn which mutations will allow fragmentation.

Scheme 16


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ACKNOWLEDGMENTS We thank the Natural Sciences and Engineering Council of Canada for support.

REFERENCES 1. 2. 3.

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R Kluger. Thiamin diphosphate: a mechanistic update on enzymic and nonenzymic catalysis of decarboxylation. Chem Rev 87:863–876, 1987. RL Schowen. In: ML Sinnott, ed. Comprehensive Biological Catalysis: A Mechanistic reference. San Diego: Academic Press, 1998. FJ Alvarez, J Ermer, G Huebner, A Schellenberger, RL Schowen. Catalytic power of pyruvate decarboxylase. Rate-limiting events and microscopic rate constants from primary carbon and secondary hydrogen isotope effects. J Am Chem Soc 113:8402–8409, 1991. R Kluger, J Chin, T Smyth. Thiamin-catalyzed decarboxylation of pyruvate. Synthesis and reactivity analysis of the central, elusive intermediate, alphalactylthiamin. J Am Chem Soc 103:884–888, 1981. LJ Dirmaier, GA Garcia, JW Kozarich, GL Kenyon. Inhibition of benzoylformate decarboxylase by [ p-(bromomethyl)benzoyl]formate. Enzyme-catalyzed modification of thiamin pyrophosphate by halide elimination and tautomerization. J Am Chem Soc 108:3149–3150, 1986. MS Hasson, A Muscate, GTM Henehan, PF Guidinger, GA Petsko, D Ringe, GL Kenyon. Purification and crystallization of benzoylformate decarboxylase. Protein Sci 4:955–959, 1995. MS Hasson, A Muscate, MJ McLeish, LS Polovnikova, JA Gerlt, GL Petsko, GA Petsko, D Ringe. The crystal structure of benzoylformate decarboxylase at 1.6-A˚ resolution: diversity of catalytic residues in thiamin diphosphate-dependent enzymes. Biochemistry 37:9918–9930, 1998. EA Sergienko, J Wang, L Polovnikova, MS Hasson, MJ McLeish, GL Jordan, F Jordan. Spectroscopic detection of transient thiamin diphosphate– bound intermediates on benzoylformate decarboxylase. Biochemistry 39: 13862–13869, 2000. R Kluger, T Smyth. Interaction of pyruvate–thiamin diphosphate adducts with pyruvate decarboxylase. Catalysis through ‘‘closed’’ transition states. J Am Chem Soc 103:1214–1216, 1981. H Stetter, G Dambkes. Uber die praparative Nutzung der Thiazoliumsalzkatalysierten Acyloin- und Benzoin-Bildung. II. Herstellung unsymmetrischer Acyloine und alpha-Diketone. Synthesis 1977:403–404, 1977. T Ugai, S Tanaka, S Dokawa. Thiamin catalysis of the benzoin condensation. J Pharm Soc Japan 63:269, 1943. R Breslow, R Kim. The thiazolium-catalyzed benzoin condensation with mild base does not involve a dimer. Tetr Lett 35:699–702, 1994. R Kluger. Lessons from thiamin-watching. Pure and Applied Chemistry 69: 1957–1967, 1997.

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14. R Breslow, C Schmuck. The mechanism of thiazolium catalysis. Tetrahedron Letters 37:8241–8242, 1996. 15. JJ Mieyal, G Bantle, RG Votaw, IA Rosner, HZ Sable. Coenzyme interactions. V. The second carbanion in reactions catalyzed by thiamin. J Biol Chem 246:5213–5219, 1971. 16. JJ Mieyal, RG Votaw, LO Krampitz, HZ Sable. Evidence for a second carbanion in the mechanism of thiamin catalysis. Biochim Biophys Acta 141:205– 208, 1967. 17. R Kluger, JF Lam, JP Pezacki, C-M Yang. Diverting thiamin from catalysis to destruction. Mechanism of fragmentation of N(1V)-methyl-2-(1-hydroxybenzyl)thiamin. J Am Chem Soc 117:11383–11389, 1995. 18. R Kluger, JF Lam, CS Kim. Decomposition of 2-(1-hydroxybenzyl)thiamin in neutral aqueous solutions: benzaldehyde and thiamin are not the products. Bioorg Chem 21:275–283, 1993. 19. R Kluger, IF Moore. Destruction of vitamin B1 by benzaldehyde. Reactivity of intermediates in the fragmentation of N1V-benzyl-2-(1-hydroxybenzyl)thiamin. J Am Chem Soc 122:6145–6150, 2000. 20. JR Keeffe, WP Jencks. Elimination reactions of N-(2-(p-nitrophenyl)ethyl) alkylammonium ions by an E1cB mechanism. J Am Chem Soc 105:265–279, 1983. 20a. LJ Reynolds, GA Garcia, JW Kozarich, GL Kenyon. Differential reactivity in benzoylformate decarboxylase. Biochemistry 27:2212–2217, 1988. 21. J Crosby, GE Lienhard. Mechanisms of thiamin-catalyzed reactions. A kinetic analysis of the decarboxylation of pyruvate by 3,4-dimethylthiazolium ion in water and ethanol. J Am Chem Soc 92:5707–5716, 1970. 22. J Crosby, R Stone, GE Lienhard. Mechanisms of thiamin-catalyzed reactions. Decarboxylation of 2-(1-carboxy-1-hydroxyethyl)-3,4-dimethylthiazolium chloride. J Am Chem Soc 92:2891–2900, 1970. 23. JA Gutowski, GE Lienhard. Transition-state analogs for thiamin pyrophosphate–dependent enzymes. J Biol Chem 251:2863–2866, 1976. 24. IF Moore, R Kluger. Substituent effects in carbon-nitrogen cleavage of thiamin derivatives. Fragmentation pathways and enzymic avoidance of cofactor destruction. J Am Chem Soc 124:1669–1673, 2002. 25. JP Guthrie. Uncatalyzed and amine-catalyzed decarboxylation of acetoacetic acid: an examination in terms of no barrier theory. Bioorg Chem 30:32–52, 2002. 26. HC Dunathan. Conformation and reaction specificity in pyridoxal phosphate enzymes. Proc Nat Acad Sci (USA) 55:712–716, 1966.

18 Structural and Functional Organization of Pyruvate Dehydrogenase Complexes Z. Hong Zhou and James K. Stoops University of Texas Health Science Center at Houston Medical School, Houston, Texas, U.S.A. Lester J. Reed University of Texas at Austin, Austin, Texas, U.S.A.

I. INTRODUCTION Pyruvate dehydrogenase complex (PDHC) belongs to a family of enzymes that catalyzes the oxidative decarboxylation of a-keto acids. It is a classic example of a multienzyme complex. As illustrated in Figure 1, PDHC catalyzes a key reaction at the junction of glycolysis and the citric acid cycle that leads to the decarboxylation of pyruvate and the acetylation of coenzyme A (CoA) by a multistep reaction involving five different cofactors: thiamine diphosphate (TPP), protein-bound lipoyl moiety (LipS2), nicotinamide adenine dinucleotide (NAD+), flavin adenine dinucleotide (FAD), and CoA (1–3). With an Mr f106–107, PDHC is the largest and one of the most complex multienzyme systems known, and the elucidation of its structural organization and functional mechanisms remains one of the most challenging problems (for reviews, see Refs. 2–6). The core of PDHC is formed by dihydrolipoamide acetyltransferase (E2), which has both functional and structural roles and serves as a scaffold to 309

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Figure 1 Reactions catalyzed by mammalian PDHC. TPP, thiamin diphosphate; LipS2 and Lip(SH)2, lipoyl moiety and its reduced form. In addition to the four major components that are present in yeast and microbes, the mammalian PDHC contains pyruvate dehydrogenase kinase (PDK) and pyruvate dehydrogenase phosphatase (PDP).

which the other components are attached (2–5). These include pyruvate dehydrogenase (E1) and dihydrolipoamide dehydrogenase (E3). E3 requires a binding protein (BP) to anchor it to the core of the yeast (7,8) and mammalian PDHC (8,9), though in Escherichia coli and Bacillus stearothermophilus, BP is not required (2–5). In mammalian PDHC, four pyruvate dehydrogenase kinase and two pyruvate dehydrogenase phosphatase isoforms are also present, and they function in controlling the activation state of the PDHC by determining the fraction of active (nonphosphorylated) E1. Adaptable control of PDHC activity is required to carry out diverse tasks in the management of fuel consumption and storage, which is achieved by the tissue and metabolic state–specific expressions and the regulatory properties of the dedicated kinases and phosphatases (10). E2 contains domains that bind the other constituents of the functional PDHC (2–5) (Fig. 2). The small binding domain of E2 of eukaryotes binds only E1, whereas the structurally related E2 domains from microbes can bind either E3 (e.g., E. coli PDHC) or both E3 and E1 (e.g., B. stearothermophilus PDHC). The C-terminal catalytic and self-association domains of eukaryotic E2 are apparently unique in binding BP. Binding protein is composed of three linker-connected domains analogous to those of E2 (Fig. 2). The N-terminal half of E2 consists of one to three lipoyl domains (L) and a small E1-binding domain, connected by Ala- and Pro-rich linkers of 20–30

3D Structures of PDHCs

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Figure 2 Diagrammatic representation of the structural domains of E2 and BP. The domains are connected by flexible linkers (depicted by wavy lines). The arrow denotes the position of truncation in the yeast E2 to generate tE2. L: lipoyl domain. The mammalian E2 is also shown for comparison. Note that an extra lipoyl domain is present in the mammalian E2. Pivot refers to the site above which the lipoyl domain rotates (see Sec. III.E and Figure 7).

residues in length (Fig. 2). The number of lipoyl domains varies in E2 from different species, for example, the yeast E2 subunit has only one L domain whereas the mammalian E2 has two: L1 and L2 (Fig. 2). It is postulated that the mobility of the linker regions allows the lipoyl domains to be delivered, through a ‘‘swinging arm’’ active-site coupling mechanism (2), to the E1, E2, and E3 components where the prosthetic group of E2 extends into active-site channels. Specific interaction of the lipoyl domain with E1 is essential for efficient E1 catalysis. Electron microscopy (11–14) and X-ray crystallography (15–17) have revealed two fundamental morphologies of the E2 cores. The cubic E2 core that exists in some microbes (such as E. coli) has 24 subunits arranged with octahedral symmetry, whereas the pentagonal dodecahedral E2 core from eukaryotes and some Gram-positive bacteria has 60 subunits arranged with icosahedral symmetry. The E2 subunits form a cone-shaped trimer at each of the 8 and 20 vertices of the cubic and dodecahedral structures, respectively. These trimers are interconnected by bridges to form a cagelike complex (13, 15–17). X-ray crystallography and NMR have been used to solve the atomic structures of several individual domains or components and the cubic form of a truncated E2 core (17–26). However, due to the inherent flexibility and un-

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precedented dynamics of the components in the PDHC (27) (see later), no atomic resolution structures have been determined for the entire PDHC or the dodecahedral form of the E2 core, except for a computationally derived model of the B. stearothermophilus tE2 core (28). In this regard, electron cryomicroscopy (cryoEM) is an appropriate and powerful method for examining protein dynamics and thus provides a unique tool to elucidate 3D structural information of the PDHC that is otherwise unavailable. Electron cryomicroscopy affords a snapshot of the molecular composition of the protein preparation at room temperature in the absence of constraints imposed by a crystal lattice with a shutter speed of f105 seconds (29,30), i.e., the time required to reduce the temperature of the specimen from room temperature to approximately 170jC, at which point molecular motion is frozen. This rapid freezing of the molecules in solution from room temperature provides a reliable means of determining the molecular structures representing that at the room temperature. We have employed both methylamine tungstate stain electron microscopy (EM) and cryoEM to elucidate the 3D structural organization of Saccharomyces cerevisiae and bovine kidney PDHC and its subcomplexes (13,27,31,32). Three-dimensional reconstruction, together with new computational methods in classifying images according to their sizes, has revealed the structural organization of the E2 core and its association with BP, E3, and E1 in a functional PDHC and their unusual structural flexibility/dynamics, which is implicated in the multifunctional roles of this remarkable complex. This chapter presents some of our latest structural data and summarizes our current understanding of the structural and functional organization of PDHC based on structural information obtained by integrating our cryoEM data with atomic structures of individual components.

II. MATERIALS AND METHODS A. Preparation of Native PDHC and Recombinant Subcomplexes The S. cerevisiae tE2 (residues 206–454 of E2), BP and E3, were overexpressed in E. coli. The BP monomers, E3 dimers, tE2, and E2 cores were purified to near homogeneity as described in Refs. 7, 27, and 33. The recombinant tE2 and E2 cores exhibited catalytic activity similar to that of wild-type E2. Purified tE2 and BP were mixed in a molar ratio of 1:24 to generate tE2/BP subcomplexes. E3 was then added to the tE2/BP preparation (molar ratio 24:1) to obtain tE2/BP/E3 subcomplexes. The mammalian PDHC was purified from bovine kidney as described in Ref. 34. Its subunit composition was


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estimated to be about 22 E1 tetramers, 60 E2 monomers, 12 BP monomers, and 6 E3 dimers (32). B. Electron Microscopy Electron cryomicroscopy of S. cerevisiae tE2, E2, tE2/BP, tE2/BP/E3, and bovine kidney PDHC were performed using established procedures (31). Briefly, a 3-AL sample (the concentration of each sample was adjusted such that the tE2 content in the sample was f0.2 mg/mL) was deposited onto a freshly prepared holey carbon EM grid, blotted, and quickly frozen in liquid ethane cooled by liquid nitrogen. Focal-pair micrographs were recorded on Kodak SO 163 film in a JEOL 1200 electron cryomicroscope operated at 100 kV with a combined electron dosage of about 18 electrons/A˚2 at 50,000 magnification. The first image in the focal pair was recorded at a targeted underfocus value of f1.0 Am and the second one at 2.5–3.0 Am underfocus. C. Computer Image Processing and Reconstruction Our experimental data include focal pairs comprising close-to- and far-fromfocus images. The former contains higher-resolution but lower-contrast image data for use in the final reconstruction at higher resolution, whereas the latter contains particle images of relatively high contrast from which a preliminary low-resolution 3D reconstruction was readily obtained (35–37). Computed projections of this initial low-resolution 3D reconstruction are reasonably noise-free and were used to assist in determining the center and orientation parameters of the corresponding close-to-focus particle images. Iterative algorithms were used to refine the center and orientation parameters, and 3D reconstructions were carried out using Fourier–Bessel synthesis methods (35–37). The structural components of interest in each map of the PDHC subcomplexes were visualized using the Iris Explorer software (NAG, Inc., Downers Grove, IL) with custom designed modules. The atomic coordinates of the crystal structure of the dodecahedral form of B. stearothermophilus tE2 (PDB identification No.: 1B5S) were kindly provided by Aevarsson and Hol (28). The atomic coordinates of the Pseudomonas putida E1 tetramer (PDB id: 1QS0) (26), the E2 lipoyl domain of B. stearothermophilus (PDB id: 1LAC) (22), and the E3-binding domain of E. coli E2 (PDB id: 1BBL; i.e., the putative E1-binding domain of E2) (38) were downloaded directly from the Protein Data Bank. The areas of interest of the structures were rendered and exported to Open Inventor format using either Ribbons (39) or Weblab ViewerPro

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(Accelryes, San Diego, CA) or directly converted to electron density maps using a Gaussian filter to resolutions similar to the cryoEM structures for further 3D comparisons.

D. Assessment of Particle Size Variation The relative sizes of PDHC or its subcomplexes were determined using the program sizeDiff (27,37). SizeDiff uses an iterative method that minimizes the Fourier cross-common-lines-phase residuals between the particle images and projections computed from a 3D model. After the orientation and center parameters of each particle were determined and the structure refined to f30-A˚ resolution, a preliminary 3D reconstruction was calculated by combining all of the refined particles. Approximately 10–15% of the particles were eliminated in this step due to poor phase residuals. This ‘‘average’’ reconstruction was then used as the model for the first round of size determination. Initially, 20 projections computed at regularly spaced orientation intervals were used in the sizeDiff analysis. Each particle image was then isotropically scaled to best match the projections by minimizing the averaged cross-common-lines-phase residual between the particle and the projections. Further classification of the particles and the refinement of the reconstruction were accomplished iteratively by utilizing the particles within F 1% of the designated size group until size difference within each group converged to within 2%.

III. RESULTS AND DISCUSSION A. CryoEM The cryoEM images of expressed S. cerevisiae tE2 (Fig. 3a) and E2 (Fig. 3b) cores embedded in vitreous ice show characteristic dodecahedral particles that appear similar despite the presence of lipoyl and E1-binding domains in the E2 (see Fig. 2). The similarity between the tE2 and E2 images shows that the N-terminal half of the E2 subunit is extended, presumably in a flexible and nonglobular configuration and, consequently, is not readily seen in the noisy cryoEM images of the E2 cores. In this regard, the image contrast of the E2 cores is notably lower than that of the tE2 cores at similar imaging conditions. This phenomenon is most likely due to the thicker ice needed to fully embed the E2 cores with the extra N-terminal domains extending from the dodecahedral core. The E1 tetramers bound to the E2 core in the native bovine kidney PDHC increased the apparent diameter of the molecule from f250 to f500 A˚ (Fig. 3c). Occasionally the images of individual PDHC particles


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Figure 3 CryoEM images of tE2 core (a), E2 core (b), and PDHC (c). The S. cerevisiae recombinant tE2 (a) and full-length E2 (b) cores were imaged at 100 kV in a JEOL 1200 electron cryomicroscope as described in Ref. 31. (c) The native bovine kidney PDHC was imaged similarly (32). One representative particle of tE2, E2, and PDHC is encircled in each micrograph.

reveal the characteristic views of the pentagonal dodecahedron-shaped core to which the E1 components are bound (Fig. 3c). B. Three-Dimensional Structures of Yeast tE2 Core and tE2 Complexed with Binding Protein The first 3D structure of the S. cerevisiae dodecahedral tE2 was determined by stain and cryoEM to about 25-A˚ resolution (13). The tE2 structure has since been improved to about 15-A˚ resolution (Fig. 4a). The trimer resolved in this improved structure shows a striking resemblance to the trimers present in the X-ray crystal structures of both the dodecahedral tE2 core from B. stearothermophilus (28) and the cubic tE2 core from Azotobacter vinelandii (17) when they were compared at similar resolution (Fig. 4b–c). In addition, the pentagonal dodecahedral scaffolds of the S. cerevisiae tE2 and E2 are not distinguishable at about 20-A˚ resolution, indicating that only the C-terminal half, i.e., the catalytic and association domain, of the E2 subunit contributes to the formation of the cagelike pentagonal dodecahedral scaffold of the PDHC core (27). The structural similarity between the scaffolds affords structural validation of numerous biochemical studies (2–5) showing that the N-ter-

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Figure 4 Shaded surface representations of the 3D structures of tE2 cores (a–c) and tE2 cores with BP bound (tE2/BP) (d–e). (a) 3D structure of the S. cerevisiae tE2 core reconstructed from electron microscopic images to about 15-A˚ resolution. (b, c) Shaded surface representations of the 3D structures of B. stearothermophilus tE2 (c) (28) and A. vinelandii cubic tE2 (d) (17) obtained by filtering their atomic models to 12-A˚ resolution using a Gaussian filter. (d) 3D reconstruction of tE2/BP obtained by cryoEM atf20-A˚ resolution. The map is displayed at a lower threshold than those in parts (a–c). (e) Same as part (d) except that a portion of the map is removed to better reveal the internal spherical density that is contributed by BP.

minal half of intact E2 is composed of flexible domains that are disordered and thus unresolved in the E2 core reconstruction obtained by averaging many particles. Difference imaging of tE2/BP and tE2 cores demonstrated that BP resides inside the tE2 scaffold (Fig. 4d–e) (31). The cryoEM structure of tE2/BP shows that the tE2 cage encloses a spherical-shaped mass density of about 100-A˚ diameter that is attributed to the bound BP molecules (Fig. 4d–e).


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C. Direct Evidence for the Size and Conformational Variability of the ‘‘Breathing’’ E2 Core and Its Functional Relationship to Protein Dynamics In the process of improving the resolution of yeast E2 core reconstructions, we identified a surprising size variation among the tE2 and E2 cores in solution, which represents an extraordinary example of protein dynamics (27). Careful visual examination of the tE2 images in cryoEM fields showed that their sizes are variable (Fig. 5b) (13,27). The possible implication and significance of these initial visual findings led to the development of an algorithm for systematically evaluating the size variability of noisy cryoEM images (27). Our classification showed that both the frozen-hydrated and stained molecules of tE2 in the same field vary in size by approximately 20% and that the distribution of the number of particles as a function of particle sizes has a bell shape (Fig. 5a). Initially we were very skeptical about the observation that a nearly homogenous preparation of tE2 molecules exhibits a 20% size variation in the same image field. However, visual inspection of cryo and stain electron micrographs of different preparations of tE2 showed significant size variation (31). In previous studies, molecules that varied more than 3% from the average size were not used in the reconstruction. In other words, the size variation was initially attributed to magnification variations and was considered a hindrance (nuisance) in computing the 3D structures. Subsequently, we investigated several trivial possibilities that could have contributed to the size variation. These included (a) magnification variation of the EM, (b) distortion of the molecules, and (c) incomplete structures, all of which have been ruled out experimentally to be the cause of the size variations observed in our reconstructions (27). With our skepticism and even disbelief allayed by these results, we documented and characterized this extraordinary size variability by determining the 3D structures of the molecules representative of the size groups from the stain and cryoEM images. The 3D reconstructions from frozenhydrated molecules in each size group are very similar and have similar resolution (20–25 A˚) (Fig. 5c–d). There is approximately a 40-A˚ difference in the diameter of the smallest and largest structures that corresponds to f14-A˚ variation in the length of the bridge connecting adjacent trimers (Fig. 5c). The variable size of the structures is not related to an isotropic change in the size of the molecule; rather, the superimposed reconstructions representative of the 0.9 and 1.1 size groups demonstrate that the size variability is related primarily to a change in distance of the trimers on their three-fold axes to the center of the molecule (Fig. 5d). As a consequence, the distance between adjacent trimers (the length of the interconnecting bridge) is also variable. We

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proposed that the size change of the molecule involves a significant contribution from a synchronous change in the length of the bridges (27). The relevance of the size variation of the tE2 core may be questioned because the tE2 core lacks nearly half of the protein of the E2 subunit. Consequently, we have determined the size variation of full-length recombinant E2 cores. A plot of the number of E2 molecules versus relative size gives the similar bell-shaped curve associated with the tE2 data sets (27). The overlay structures of the 0.95 and 1.05 E2 reconstructions are entirely consistent with those corresponding to tE2. Thus, we conclude that the size variability is a fundamental property of the C-terminal catalytic and selfassociation domain of the E2 subunit. The X-ray structure of B. stearothermophilus tE2 offers a plausible explanation for the flexible springlike connectivity between adjacent trimers

Figure 5 Size distribution of the recombinant S. cerevisiae tE2 core. (a) The bar graph shows the approximate relative size distribution of about half of the 3940 particular images of the frozen-hydrated tE2 cores. The bell-shaped distribution profiles indicate that there is a continual variation in the size of the molecules in the preparation. (b) A comparison of selected images from the different size groups shown in part (a). The circle around the images corresponds to the diameter of the images from the 1.1 size group and serves as an aid in determining their relative sizes. (c) Shaded surface views of the tE2 structure reconstructed from three selected size groups. (d) Superposition of the 3D structures reconstructed from particles in the 0.90 (semitransparent surface representation) and 1.10 (wireframe) size groups. The reconstructions are rendered at the same threshold (contour level). The size change is related to a variation of the distance of the trimers along their threefold axes from the center of the molecule. Stain images recorded at room temperature gave a similar size distribution, thus supporting the proposal that cryoEM gives a snapshot of the molecular distribution at room temperature (27). (e) Blown-up view of the 3D structure of B. stearothermophilus tE2 along an icosahedral twofold axis. The ribbon representation of the bridge on its twofold axis shows that there is an f10-A˚gap between adjacent trimers and that the trimers are interconnected by only two C-terminal extensions of the polypeptide chain from the opposing subunits. (f ) Ribbon diagram of the residues from the C-terminal region of the B. stearothermophilus tE2 that comprise the putative spring that interconnects adjacent trimers in the dodecahedron tE2 core. A loop region comprising residues 397–403 is anchored to a h-sheet. This is followed by a four-turn a-helix, beginning with residue 403 and disrupted by Pro-420, that directs residues 421–425 to the subunit of the adjacent trimer. The C-terminus resides in a hydrophobic pocket (HP) of the adjacent subunit to form the ball-and-socket connection. Residues 403–425 form a cantilever-like structure, which is attached to the loop at its N-terminus. The hydrophobic pocket readily accommodates the rotation of the C-terminal Met during expansion and contraction of the core.


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(Fig. 5e–f ). The X-ray structure of B. stearothermophilus tE2 shows that there is an f10-A˚ gap between adjacent trimers and that the trimers are interconnected by the potentially flexible C-terminal ends of two adjacent subunits (Fig. 5e). We proposed that this springlike feature is involved in a thermally driven expansion and contraction of the core and, since it appears to be a common feature in the phylogeny of PDHCs, that protein dynamics is an integral component of the function of these multienzyme complexes. Although the subunits of the trimers exhibit extensive connections between them, adjacent trimers are held together only by interactions of the C-terminal methionine of one subunit with the hydrophobic pocket of the adjacent subunit (Fig. 5e). The X-ray structure also suggests a springlike connection between adjacent trimers consisting of a C-terminal loop (residues 397–402) that is anchored to a h-sheet followed by an a-helix (residues 403–419) (Fig. 5f). Proline 420 at the C-terminal end of the rod-shaped a-helix introduces an elbow bend that directs the C-terminal methionine into the hydrophobic pocket. We believe that the E2 cores are exceptional examples of what has been designated ‘‘soft proteins’’ (40). The association energy due to hydrophobic interactions is similar to thermal energy at room temperature, 1 kcal/mole (41). It is noteworthy that B. stearothermophilus, Enterococcus faecalis, S. cerevisiae, and human dodecahedral E2 molecules have a proline residue that is five to eight residues from their C-termini, and this residue is often associated with a flexible region in the polypeptide chains (42). Furthermore, there is a 55% sequence homology between the C terminal 29 residues of the E2 molecules from these species, indicating that they all share the loop region, cantilever, and ball-and-socket feature seen in the X-ray structures of tE2 from B. stearothermophilus and E. faecalis and thus exhibit similar flexibility. This conservation of the sequence homology through all phylogeny indicates that the flexibility associated with the yeast PDHC has an important role in its function. Our comparisons of the various size structures showed that there are conformation changes in the bridge between trimers and the trimers themselves (27). Moreover, the X-ray structure shows that there is a 29-A˚-long channel through which CoA passes from its binding site inside of the core to the outer surface location of the lipoamide-binding site (28). In conjunction with the protein dynamics of the core, this channel may contract and open in a pumping action to promote the movement of CoA through this channel. The breathing core may augment the movements of the lipoyl domain swinging arms between the catalytic centers (see upcoming Sec. III.E), and the apparent movement of the entire complex may augment substrate channeling and promote catalysis by mechanisms that are just beginning to be understood (43,44). We have also shown that protein dynamics is an integral part of the structural organization of the fully assembled PDHC (27). In this study, the


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3D reconstructions of E2 with BP, E3, and E1 bound showed that the inner core and the outer ring of protein density corresponding to E1 display a variable distance from the center of the center of the core (45). The realization that significant size variation is an integral property of tE2, E2, and the fully assembled PDHC has led to a change in our data-processing approach of these complexes. Consequently, we have gained major new insights into the structure–function relationships of one of the most challenging problems in structural biology—the elucidation of the structural organization of one of the most complex multienzyme systems known. D. Size Variation Classification is Crucial to the Localization of E3 in tE2/BP/E3 Complex We have also improved the structure of the yeast tE2/BP/E3 complex in an attempt to better visualize the sites of contact between tE2 and E3 (Fig. 6). Initial tE2/BP/E3 reconstructions, which were obtained without considering their possible size variations, did not reveal structural features significantly different from those seen in tE2/BP (Fig. 4d–e). Our first approach to this problem was to saturate the tE2/BP with E3 molecules (E2/BP:E3 molar ratio = 1:24). However, 3D reconstruction of the tE2/BP/E3 images did not reveal significant differences from that of the tE2/BP complex unless the particles were classified into different groups according to the relative sizes and then independently reconstructed, as illustrated by the two structures independently reconstructed from the size groups of 0.94–0.96 and 1.04–1.06 (Fig. 6c). Reconstruction from the particles in the smaller size group revealed pentagonal openings similar to those seen in the tE2/BP complex (see Fig. 4d–e). However, the reconstruction from particles in the larger size group reveals an extra density almost complete blocking the pentagonal openings. The 12 pentagon-shaped densities nearly fill the pentagonal openings on each face of the tE2 dodecahedron (Fig. 6b). Because this density is almost completely absent from the structure of tE2/BP complex (Fig. 4d–e), we attribute the bulk of this density to a bound E3 dimer. Therefore, contrary to previous belief, the 12 large openings in the yeast dodecahedral E2 core permit the entrance of E3 and BP into the central cavity to interact with the interior of the E2 component (31). It should be pointed out that, although we have definitely located E3 dimer in the E2/BP/E3 subcomplex, the E3 density revealed in these reconstructions was smeared out and distorted due to icosahedral averaging imposed during the 3D reconstruction, regardless whether only 12 E3, as presented previously (31), or 24 E3 dimers per tE2/BP, as used in the current study (Fig. 6b), were present. The reconstruction from the smaller size group did not reveal the E3 components, most likely due to a lower E3 occupancy in these particles (Fig. 6c). It is conceivable that, due to geometric constraints

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Figure 6 Localization of E3. (a) CryoEM micrograph of S. cerevisiae tE2/BP/E3 subcomplex. (b) Shaded surface view along a threefold axis of the 3D structure of tE2/BP/E3 reconstructed from a subset of particles with relatively large sizes (size group 1.03–1.06). A pinlike density fills each of the 12 pentagonal openings of the tE2/BP complex (Fig. 4d) and is attributed to an E3 dimer. (c) Classification of tE2/ BP/E3 particles according to their relative sizes. The insets show the cutaway views of the 3D reconstructions from the 0.94–0.96 size group and 1.03–1.06 size group. In the superposition of the two structures, the larger size particle is shown semitransparently.


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arising from the reduced size, the smaller E2 scaffold could not accommodate all of the 12 E3 dimers associated with each larger core. E. Structure and Functional Organization of the Native PDHC We have also determined the 3D structure of the intact bovine kidney PDHC (32) (Fig. 7). Due to the flexibility of the intermolecular interactions among its components, intact PDHCs are sensitive to sample preparation procedures. The E1-binding domains are linked to the E2 core by Pro-rich, potentially flexible linkers (see Fig. 2), and thus the deposition of the E1 tetramers could be particularly sensitive to these procedures. Our experience indicates that it is important to use freshly prepared PDHC sample and avoid freezing and thawing the sample, to prevent E1 molecules from dissociating from E2 core. As judged from the very smooth amorphous ice background in the cryoEM images of the hydrated intact PDHC, E1 molecules did not dissociate significantly from the intact PDHC during our cryoEM preparation. The 3D structure reveals that five E1 tetramers form a crown (Fig. 7a), which is linked to five E2 molecules of the pentagonal dodecahedron E2 core (Fig. 7b). Considerable effort was made to reveal the connections (linkers) that bind E1 molecules to the E2 core, since this structure would be expected to provide information that is fundamental to the appropriate docking of the E1 X-ray structure in the electron microscopy envelope (see later). Initially, all our reconstructions of the S. cerevisiae and the bovine kidney PDHCs displayed a shell of protein density surrounding the underlying core without revealing the associated linkers. The utilization of size variation analysis to classify the images according to size (27) and the utilization of closer-to-focus images was an essential step in our effort to align the images with enough precision to reveal the connections. The Fresnel fringe effect introduces significant negative contrast around the core, masking this component of the reconstruction at the larger defocus values used previously. The bovine kidney native PDHC structure shows that potentially 60 units for acetyl-CoA synthesis are organized in sets of three at each of the 20 vertices of the dodecahedral core (Fig. 7e–f). Consequently, the structural and functional organization of the dodecahedral PDHC consists of three E1 molecules bound to one E2 trimer (Fig. 7e–h). We propose that each E2 trimer and its three connected E1 dimers, together with its adjacent E3 molecule in the pentagonal opening of the scaffold, comprise the functional unit of the scaffold (Fig. 7e–h). Consequently, our structure provides a resolution of the longstanding question regarding what constitutes the ‘‘swinging arm’’ and its length (6). The E1-binding domain of E2 serves as a pivot or anchoring point for a swinging arm comprising the outer linker and

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the two lipoyl domains (Fig. 7h). The swinging arm rotates about the E1binding domain of E2, which is centrally locatedf50 A˚ from the E1, E2, and E3 active sites (Fig. 7h). The three linkers that surround the E2 trimer form a cage that is covered by accompanying E1 molecules. This cage may function as a shelter that is important in ensuring the successful transfer of the acetylated dihydrolipoyl moiety to the acetyltransferase site that resides f95 A˚ below the E1 catalytic site (Fig. 7f, h). Because one E1 active site faces the inside of this cage, whereas the other faces the outside (Fig. 7h), this arrangement may favor the former in the catalytic sequence and thus provide a structural basis for the half-of-site reactivity exhibited by E1 (6). The appearance of the inner linker in the structure and the f40-A˚ resolution corresponding to the E1 shell shows that flexibility of E1 and its linker is largely constrained. It seems paradoxical that the E1 components are constrained by the inner linker, since this tether does not appear in the reconstruction of E2, due to its flexibility. We propose that the E1 molecules exhibit weak but significant interaction with each other; consequently, they

Figure 7 3D reconstruction of bovine kidney PDHC at 30-A˚ resolution and fitting of atomic structures of individual components. (a) Shaded surface view along an icosahedral threefold axis. Icosahedral five-, three-, and twofold axes are indicated. (b) The top half of the structure was removed to reveal the inner linker attached to the underlying core. The BP/E3 components associated with the core are not revealed, probably due to their low occupancy in this preparation (less than six E3 dimers bound). The inner linker isf50 A˚ in length and serves to attach E1 to the E2 scaffold. (c) The E1 densities were removed to reveal the underlying E2 core and its inner linkers. (d) Zoom-in view of the region highlighted by the dotted line in part (c) superimposed with the atomic structure (ribbons) of B. stearothermophilus tE2. The cryoEM and atomic structures were aligned by matching their icosahedral five-, three- and twofold axes. The inner linker densities are directly opposite the N-terminal helix (H1) of the atomic structure. (e) Top view of the cutaway structure comprising an E2 trimer and its associated three E1 tetramers. (f ) Side view of part (e), showing the three inner linkers that bind three E1 tetramers to the core. (g, h) Same as parts (e) and (f ), except shown as wireframe representation and that the atomic structures (shown as ribbons) of P. putida E1 (26) and two copies of the B. stearothermophilus lipoyl domain (22) [denoted by L and the light arrows in part (h)] and the putative E1-binding domain (38) are docked in the cryoEM structure envelope. The light gray arrow denotes the E1 active site, which faces toward the center of the cagelike structure. The single lipoyl domain of E2 appears to bind to this site in preference to the E1 active site facing to the outside of this functional unit. The rods denote the crystallographic twofold and threefold axes of E1 and E2, respectively. The E1-binding site (indicated by the dark arrow) on the E2 inner linker is centrally locatedf50 A˚ from the E1, E2, and E3 active sites and serves as the pivot about which the swinging arm rotates.


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are grouped together in clusters above the underlying trimers. This interaction may provide positional stability to the organization of the E1 molecules. This interaction would be favored at the high local concentration of the E1 molecules (about 100 mg/mL). Consequently, in an attempt to stabilize the E1 molecules about the core and to eliminate their down-weighting in the cryoEM map due to their low occupancy in the native bovine kidney PDHC preparation (about 22 E1 tetramers bound to each E2 dodecahedron), we determined the structure of S. cerevisiae PDHC using a preparation with nearly saturated E1 (f60 E1 tetramers bound) (45). Surprisingly, the deposition of the E1 molecules around the outside of the core in this high-E1-occupancy PDHC is greatly altered from the deposition of E1 molecules as revealed in the bovine kidney PDHC (Fig. 7). The icosahedral arrangement of the E1 components is lost, as indicated by the very low Fourier shell correlation coefficient between independent reconstructions within the region of the E1 densities; therefore, we were unable to meaningfully dock the X-ray structure of E1 in the outer shell of this PDHC structure. Despite the poor resolution of the outer shell, the overall resolution of the complex reaches about 30 A˚ due to the excellent icosahedral symmetry of the inner core. Interestingly, though its density is significantly weaker, the inner linker revealed in this reconstruction (45) is located in the same position as, and has a very similar morphology to, that shown in the bovine kidney PDHC (Fig. 7b). Therefore, it appears likely that, in addition to size variation intrinsic to PDHC particles and the Fresnel fringe effect resulting from the use of large underfocus values during imaging, other factors, such as chemical differences inherent in the PDHCs from different species and/or differences in sample preparation and buffer conditions, may also be related to the robustness of the density of the inner linker. The most amazing result of this enormous change in the E1 deposition is that the PDHC specific activity is not significantly affected upon nearly saturating the E1-binding domains in this preparation. This appears to contradict the dogma that enzyme active sites have very strict requirements regarding their spatial organization for proficient catalysis. Undoubtedly, the protein dynamics of PDHC makes it possible to function appropriately even when one of its components (E1) undergoes a major rearrangement.

IV. CONCLUSIONS AND FUTURE PERSPECTIVES Electron cryomicroscopy has provided a useful tool in establishing the molecular architecture of the entire PDHC and in demonstrating the unusual property of structural dynamics of this multienzyme complex and its subcomplexes. Difference imaging using PDHC and recombinant subcomplexes has allowed the localization of the functional components associated with the


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pentagonal dodecahedral scaffold formed by E2, including E1, BP, and E3. The localization of these components in the cryoEM structure of the entire PDHC and the fitting of the atomic structures of individual functional domains into the cryoEM structures have provided considerable insights into the mechanism by which the series of reactions are catalyzed by this multienzyme complex. Future studies toward higher resolution by cryoEM will further delineate the significance of the ‘‘breathing’’ molecules in promoting their functions. For example, the imaging of the cantilever-like structure in the 0.9 and 1.1 size structures at 8 to 9-A˚ resolution may make it possible to document the mechanism for the largest size variation of a macromolecule known to date and to verify the significance of the ball-and-socket connection near the twofold bridge in the protein dynamics of the E2 core. Such studies will also determine the role of other conformational changes that are important for the function of the PDHC. The 3D reconstruction of mammalian PDHC bound with kinases and phosphatases may offer new insights into the mechanisms of its regulatory control, which is essential for its function in cellular metabolism.

ACKNOWLEDGMENTS The research activities described here have been supported in part by grants from the American Heart Association (0240216N to ZHZ) and NIH (AI46420 and CA94809 ZHZ and EB00276 & HL42886 to JKS), the Welch Foundation (AU-1492 to ZHZ) and the March of Dimes Birth Defects Foundation (5-FY99-852 to ZHZ). ZHZ is a Pew Scholar in Biomedical Sciences. We thank Steven Kolodziej for assistance in preparing Figures 4d and 6a. REFERENCES 1. 2. 3. 4.

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19 The Pyruvate Dehydrogenase Multienzyme Complex Richard N. Perham Cambridge University, Cambridge, England Jacqueline S. Milne and Sriram Subramaniam National Institutes of Health, Bethesda, Maryland, U.S.A.

I. INTRODUCTION The pyruvate (PDH), 2-oxoglutarate (OGDH), and branched-chain 2-oxo acid (BCDH) dehydrogenase complexes are members of a family of related enzymes that catalyze the oxidative decarboxylation of particular 2-oxo acids, generating the relevant acyl-coenzyme A and NADH as the principal products. They occupy key positions in metabolism: at the junction of glycolysis and the citric acid cycle, in the citric acid cycle itself, and in the metabolism of the branched-chain amino acids, leucine, isoleucine, and valine. In the PDH complex, the three component enzymes are pyruvate decarboxylase (E1p; EC 1.2.4.1), dihydrolipoyl acetyltransferase (E2p; EC 2.3.1.12), and dihydrolipoyl dehydrogenase (E3; EC 1.8.1.4). E1 catalyzes the initial decarboxylation, utilizing thiamine diphosphate (TDP) as a cofactor, and the subsequent reductive acetylation of a lipoyl group covalently attached to a lysine residue in E2. E2 catalyzes the transfer of the acyl group to CoA; and E3 concludes the process by reoxidizing the dihydrolipoyl group and regenerating its dithiolane ring with the concomitant reduction of NAD+. Corresponding enzymes make up the OGDH and BCDH complexes; in each case, E1 and E2 are 331

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specific to the particular 2-oxo acid undergoing decarboxylation. E3, which carries out an identical reaction in each complex, is normally the same enzyme in all three instances (for recent reviews, see Refs. 1 and 2). In the PDH complex from Bacillus stearothermophilus and most Grampositive bacteria, the E2p core comprises 60 E2p chains arranged with icosahedral symmetry, whereas in the PDH complex from Escherichia coli and most Gram-negative bacteria, E2p contains 24 polypeptide chains in octahedral symmetry (1–4). Multiple copies of the E1 and E3 components (E1 generally in much higher abundance than E3) are bound tightly but noncovalently in peripheral positions around the E2 core. The OGDH and BCDH complexes follow the same structural pattern, with E2 cores (E2o and E2b) of octahedral symmetry. Thus, the intact complexes are of enormous size, with molecular masses of 5–10 106 Da and diameters of up to 50 nm, significantly bigger than a ribosome. The PDH complex of B. stearothermophilus is the one about which we know most in terms of structure and molecular mechanism. The structure of the E2 component has been elucidated by recognizing first that the E2 chain is a multidomain-and-linker structure and then solving the structures of the individual domains in turn. The lipoyl group is covalently attached in an amide linkage to the N6-amino group of a specific lysine residue of an independently folded domain (about 80 residues) that forms the N-terminal part of the E2 chain. The solution structure of the domain, determined by means of nuclear magnetic resonance (NMR) spectroscopy, revealed it as a h-barrel formed from two four-stranded h-sheets, arranged with a twofold axis of quasisymmetry. The lipoyl-lysine is located at the tip of a tight, type I h-turn protruding from one h-sheet, and the N- and C-termini are close in space on the other h-sheet, on the opposite side of the domain (5). E1 and E3 are bound to E2 by means of their interaction with another domain, this being one of only 35 amino acids, the structure of which on its own (6) and in association with the dimeric E3 (7) has also been determined. The binding of E1 and E3 to this peripheral subunit-binding domain (PSBD) is tight but noncovalent; it is also mutually exclusive (8,9), engendering the possibility of multiple structural isomers occurring in the assembly of the intact complex (10). The C-terminal domain of E2 is much larger than the other two, approximately 28 kDa, and houses the acyltransferase active site; moreover, it associates with octahedral symmetry to form the 60-mer inner core of the intact PDH complex. Its structure was solved by X-ray crystallography (11), which enabled a schematic molecular model of the whole assembly to be proposed (Fig. 1). It should be noted that thus far there is no crystal structure for the B. stearothermophilus E1 component, but a plausible model can be built by homology from the crystal structures of the highly similar heterotetrameric (a2h2) E1s of the Pseudomonas putida (12) and human (13) BCDH

PDH Complex as a Molecular Machine

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Figure 1 Schematic representation of the PDH complex of B. stearothermophilus. Each E2 chain is a domain-and-linker structure: from the N-terminus, a lipoyl domain (LD), a peripheral subunit-binding domain (PSBD), and an acetyltransferase domain, joined by extended flexible linker regions. The 60-mer E2 core is made up of 20 trimers of E2 chains. The structure is viewed from the fivefold face of the E2 core, but only one E2 trimer is shown in full, for clarity. On the upper E2 chain in this trimer, the PSBD is shown binding an E3 component, and on the lower chain it is shown binding an E1 component; the middle chain is shown naked, again for clarity. (From Ref. 1.)

complexes. It is this structure that is used in Figure 1, with E1 shown as bound to the PSBD of E2 in a 1:1 stoichiometry and close to its twofold axis, as suggested by the biochemical evidence (8). II. ACTIVE-SITE COUPLING AND SUBSTRATE CHANNELING The number of lipoyl domains per E2 chain can vary, e.g., from one in the E2p chain of B. stearothermophilus PDH complex and the E2o chain of E. coli OGDH complex, to three in the E2p chain of E. coli PDH complex (3,4). The lipoyl domain plays a vital role in coupling the reactions within the complex in

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an organized and specific manner. Acting as a ‘‘swinging arm’’ (14), the lipoyl-lysine group visits each of the active sites contributed by the three different enzymes in the complex, carried by the lipoyl domain, which is itself rendered mobile by virtue of the conformational flexibility in the segment of polypeptide chain that links it to the inner E2 core of the complex (1,4). Exact positioning of the target lysine within the protruding h-turn is essential for correct posttranslational modification by the lipoylating system(s) in vivo and in vitro (15). As shown first with the PDH complex of E. coli, free lipoic acid is a surprisingly poor substrate for E1p but is acted upon rapidly by E2p and E3. However, the lipoyl domain from E2p is an excellent substrate for E1p (kcat/ Km raised by a factor of 104). Another important feature is that the lipoyl domains from the PDH and OGDH complexes of E. coli function as substrates only for their natural partner E1s (16,17 and references therein). Similar results have been reported for the lipoyl domains of the PDH and OGDH complexes of Azotobacter vinelandii (18). Thus, the true substrate in these complexes is not lipoic acid or even lipoyl-lysine but the lipoylated domain of the E2 chain. This is the molecular basis of substrate channeling, whereby reductive acylation is confined to a lipoyl group covalently attached to a specific lysine residue of the intended E2 component (1,4). Note that, as in a conventional metabolic pathway, it is this commitment at the first enzyme-catalyzed reaction that dictates the course of the subsequent flow of substrate.

III. INTERACTION OF THE LIPOYL DOMAIN WITH E1 AND THE OVERALL REACTION MECHANISM In the crystal structures of the heterotetrameric (a2h2) E1s from P. putida (12) and human (13) BCDH complexes, the TDP in the active site lies buried at the bottom of a 2-nm-deep funnel-shaped hole at the interface between the a- and h-subunits (Fig. 2). This is well beyond the reach (1.4 nm) of the fully extended lipoyl-lysine swinging arm. A prominent surface loop in the lipoyl domain, linking h-strands 1 and 2, is present only in the h-sheet that contains the lipoyl-lysine residue and lies close in space to the lipoyl-lysine h-turn (Fig. 3A). NMR and directed mutagenesis experiments with the homologous E1p of B. stearothermophilus E1p have indicated that this loop and certain residues flanking the lipoyl-lysine residue in its h-turn are likely to make transient contact with E1 during catalysis and to be of critical importance in directing the interaction (19). Further NMR experiments make it clear that the incoming lipoyl domain makes transient contact with both E1a and E1h subunits in B. stearothermophilus E1p, consistent with the active site lying


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Figure 2 Model of the lipoyl-lysine residue of a lipoyl domain being inserted into an active site of the E1 component. Note the full extension of the swinging arm and the close apposition of the lipoyl domain to E1, which is required if the dithiolane ring is to reach the thiamine for reductive acetylation to take place. (From Ref. 12.)

Figure 3 Map of the interaction sites of the lipoyl domain of the PDH complex with its cognate E1, revealed by multidimensional NMR spectroscopy. (A) A model of the lipoyl domain from E. coli E2, illustrating the point of attachment of the lipoyl group to Lys41 in the protruding h-turn and the nearby surface loop. (B) Points of transient contact between the lipoyl domain of E2 and the E1 of the E. coli PDH complex, as indicated by chemical shift changes. (C) Points of transient contact between the lipoyl domain of E2 and the E1 of the E. coli PDH complex, as indicated by T2 changes. (From Ref. 22.)

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between them, and that other points of contact, physically remote from the lipoyl-lysine residue, are distributed over the surface of the lipoyl domain (20). In the recent crystal structure of the dimeric (a2) E1p from the E. coli PDH complex (21), the TDP is again found lying at the bottom of a long funnel-shaped entrance to the active site, suggesting that a similar interaction with the relevant E. coli lipoyl domain must apply. This too is consistent with NMR experiments that indicate that a significant number of residues on the lipoyl domain, largely restricted to the lipoyl-lysine-containing half and dominated by the lipoyl-lysine h-turn, come into contact with E1p (Fig. 3B and C). No interaction between the E1 component from the OGDH complex and the apo domain from E. coli E2p could be detected, but a very weak interaction with the holo domain was observed, presumably reflecting recognition of the lipoyl lysine residue (22). To generalize, then, it would appear that the protruding h-turn housing the lipoyl lysine residue in a lipoyl domain must enter the active-site funnel in E1, bringing the nearby surface loop, among other parts of the lipoyl domain, into close contact with E1; but reductive acylation of the dithiolane ring will ensue only if the domain to which the lipoyl group is attached is also specifically recognized by E1 (22).

Figure 4 Schematic reaction mechanism of the PDH complex. Note that the lipoyl group must be attached to the cognate lipoyl domain, which is recognized by the partner E1, if reductive acetylation is to occur. (From Refs. 1 and 24.)


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The reaction mechanisms of each of the component enzymes have been much studied (1). Of particular interest here is that the acyltransferase active site in E2 closely resembles that of chloramphenicol acetyltransferase, which in turn implies that the lipoyl domain approaches the active site from the ‘‘outside’’ of the cubic assembly, whereas the CoA must enter and the acetylCoA must exit from the ‘‘inside’’ of the assembly (1,23). It is likely that the very large holes on the fourfold (of the 24 mer) and the fivefold (of the 60 mer) axes of the E2 assemblies provide the entry and exit points (see Fig. 1). The topological importance of this will become clearer later. Our understanding of the overall reaction mechanism of the complex has been boosted by an elegant study of substrate analogs, which has identified the need for a suitably placed donor in the active site of E1 to provide a proton to a sulphur atom in the dithiolane ring of the lipoyl group on E2 to facilitate the reductive acylation (24). The possible identity of this proton donor as a histidine residue was recognized (12) in the description of the crystal structure of the E1 of P. putida BCDH (see Fig. 2). The overall mechanism can thus be schematized as in Figure 4, based on that of Pan and Jordan (24), but emphasizing the need to present the lipoyl group as part of the lipoyl domain for efficacious reductive acylation to occur (as outlined earlier). IV. THE OVERALL MOLECULAR ARCHITECTURE The structure of an assembled PDH complex has thus far eluded all attacks by means of X-ray crystallography (probably because of the innate flexibility in the E2 chains and lipoyl domains and also perhaps because of structural isomerism and a lack of structural uniformity, as referred to earlier). Fortunately, cryoelectron microscopy (cryo-EM) has come to the rescue. To avoid any difficulties with structural heterogeneity, the structure of the B. stearothermophilus PDH complex has been approached by systematically assembling it from its constituent parts in vitro (25). The overall structure of the E1E2 subcomplex, in which all 60 binding sites on E2 are occupied by E1 heterotetramers, was reconstructed from multiple cryo-EM images, as shown in Figure 5. Far from the lipoyl-domains interdigitating between the E1 and E3 subunits, as hitherto supposed (1), it turns out that the E1 subunits form a shell, concentric with the icosahedral E2 inner core of acetyltransferase domains and rather uniformly placed some 9 nm above the surface of the inner core. The linker region of some 30 amino acids between the PSBD and acetyltransferase domain evidently crosses the gap only once (and is too faint to be seen in the electron density map), and the lipoyl domains, tethered to the PSBD by a linker region of some 45 amino acids, must occupy the annular space between the E1 and the E2 inner core (25).

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Figure 5 Representation of the electron density map for an E1E2 subcomplex. The overall diameter is some 475 A˚, and the E1 subunits are located rather evenly at a distance of about 90 A˚ above the surface of the E2 inner core (cf Fig. 1). (From Ref. 25.)

It is clear that a similar situation obtains with the E2E3 subcomplex, but it remains to be determined how E1 and E3 cohabit on the surface of the E2 core (JS Milne, S Subramaniam, RN Perham, unpublished work). Similar work has been reported on the bovine kidney PDH complex, but in this instance it has been claimed that the linker regions crossing the annular space between the acetyltransferase core and the E1 subunits can be identified in the electron density map and that the lipoyl domains are localized in the E1 active sites (26). Various other differences from the work of Milne et al. (25) remain to be resolved, some of which may be due to the different sources of the complexes and the fact that the E3 subunits in the eukaryotic PDH complex are bound to another sort of polypeptide chain in the E2 core, which has no counterpart in the B. stearothermophilus PDH complex. Nonetheless, the overall similarity is striking, and it seems reasonable to infer that the basic concept of an inner E2 core, an outer shell of E1, and perhaps E3 subunits, and the


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reactions involving the lipoyl domain being confined to the annular space thus created, will be common to all 2-oxo acid dehydrogenase complexes (25). V. IMPLICATIONS FOR CATALYSIS The new structural information goes a long way toward providing a molecular basis for some intriguing features of the reactions catalyzed by 2-oxo acid dehydrogenase complexes. First, constraining the lipoyl domains to operate in the annular space between the inner core of E2 acetyltransferase domains and the outer shell of E1 and E3 subunits means that the local concentration of lipoyl domains is high (perhaps of the order of 1 mM). It is difficult to estimate this with certainty without knowing more about the constraints imposed by the linker regions that tether them to the PSBDs, but it must be well above the Km for the reductive acylation catalyzed by E1, estimated to be something like 20 AM (16). It should be recalled that the true substrate for E1 is the lipoyl domain, which will diffuse much more slowly than free lipoic acid. Thus the rate enhancement by a factor of 100 or more that accompanies the covalent attachment of the lipoyl domains to E2 (1,18) is readily explained. A crucial feature of our model of the E1E2 complex (25) is that the lipoyl domain of a particular E2 chain is capable of reaching multiple E2 active sites: the three E2 active sites in the trimer directly below the relevant PSBD (within f105 A˚) and a further three neighboring E2 sites (within f140 A˚ of the same PSBD). Moreover, one of the active sites of each of six E1 heterotetramers is within f120 A˚ of the same PSBD, and an additional three E1 active sites are located within a distance of f140 A˚, making a total of nine potentially accessible to a given lipoyl domain. This may be a conservative estimate because it omits the extra range permitted by the length of the lipoyl domain itself and possible alternative fits to the electron density map. Such a structure obviates any need for a strict stoichiometric relation between the numbers of E1, E2, and E3 catalytic sites, as observed (1–4). Another longstanding conundrum is the observation that gradual excision of lipoyl domains by proteolysis or genetic manipulation is not accompanied by corresponding decreases in overall PDH complex activity (3,4) and, further, that a single E1 molecule on an E2 core can catalyze the reductive acetylation of many, if not all, of the lipoyl domains in the E2 core (27–29). Moreover, a PDH complex with optimal activity contains disproportionately few E3 molecules, for example, f42–48 E1 and f6–12 E3 for each E2 icosahedral core (10). Given that E1 catalyzes the rate limiting reaction in the complex (30,31), the multiplicity of interactions of a single lipoyl domain with different E1 and E3 molecules on the surface of the same E2 core that we propose goes a long way to providing a structural explanation for these highly unusual properties.

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It may well be that in addition there is a system of rapid intramolecular transacetylation reactions that conveys acyl groups between neighboring lipoyl domains and that would amplify the connectedness in the complex (1,4). Likewise, there is some evidence that the lipoyl group may adopt preferred orientations on the surface of the lipoyl domain (32) and that this may be accompanied by preferred trajectories of the lipoyl domains in their approach to E1 active sites (33). Such possibilities remain to be investigated further and to be put in a structural context. What is now abundantly clear is that the 2-oxo acid dehydrogenase complexes rely heavily for their function on the architectural and mechanical properties that their protein components provide and that they require a new perception of them as multifunctional catalytic machines with important lessons to teach us about other systems that depend on swinging arms in multistep catalysis (1). ACKNOWLEDGMENTS We are grateful to the Biotechnology and Biological Sciences Research Council, the Cambridge Overseas Trust, the Medical Research Council, St. John’s College, Cambridge, and The Wellcome Trust for their support at various stages of this work. We thank Mr. F. Northrop for skilled technical assistance and numerous colleagues for help and discussion. REFERENCES 1.

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RN Perham. Swinging arms and swinging domains in multifunctional enzymes: catalytic machines for multistep reactions. Annu Rev Biochem 69:961–1004, 2000. A de Kok, AF Hengeveld, A Martin, AH Westphal. The pyruvate dehydrogenase multi-enzyme complex from Gram-negative bacteria. Biochim Biophys Acta 1385:353–366, 1998. LJ Reed, ML Hackert. Structure–function relationships in dihydrolipoamide acyltransferases. J Biol Chem 265:8971–8974, 1990. RN Perham. Domains, motifs, and linkers in 2-oxo acid dehydrogenase multienzyme complexes: a paradigm in the design of a multifunctional protein. Biochemistry 30:8501–8512, 1991. F Dardel, AL Davis, ED Laue, RN Perham. Three-dimensional structure of the lipoyl domain from Bacillus stearothermophilus pyruvate dehydrogenase multienzyme complex. J Mol Biol 229:1037–1048, 1993. YN Kalia, SM Brocklehurst, DS Hipps, E Appella, K Sakaguchi, RN Perham. The high-resolution structure of the peripheral subunit-binding domain of dihydrolipoamide acetyltransferase from the pyruvate dehydrogenase multienzyme complex of Bacillus stearothermophilus. J Mol Biol 230:323–341, 1993. SS Mande, S Sarfaty, MD Allen, RN Perham, WGJ Hol. Protein–protein


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21. P Arjunan, N Nemeria, A Brunskill, K Chandrasekhar, M Sax, Y Yan, F Jordan, JR Guest, W Furey. Structure of the pyruvate dehydrogenase multienzyme complex E1 component from Escherichia coli at 1.85-A˚ resolution. Biochemistry 41:5213–5221, 2002. 22. DD Jones, KM Stott, PA Reche, RN Perham. Recognition of the lipoyl domain is the ultimate determinant of substrate channelling in the pyruvate dehydrogenase multienzyme complex. J Mol Biol 305:49–60, 2001. 23. JR Guest. Functional implications of structural homologies between chloramphenicol acetyltransferase and dihydrolipoamide acetyltransferase. FEMS Microbiol Lett 44:417–422, 1987. 24. K Pan, F Jordan. D,L-S-Methyllipoic acid methyl ester, a kinetically viable model for S-protonated lipoic acid as the oxidizing agent in reductive acyl transfers catalyzed by the 2-oxoacid dehydrogenase multienzyme complexes. Biochemistry 37:1357–1364, 1998. 25. JLS Milne, D Shi, PB Rosenthal, JS Sunshine, GJ Domingo, X Wu, BR Brooks, RN Perham, R Henderson, S Subramaniam. Molecular architecture and mechanism of an icosahedral pyruvate dehydrogenase complex: a multifunctional catalytic machine. EMBO J 21:1–12, 2002. 26. H Zhou, DB McCarthy, CM O’Connor, LJ Reed, JK Stoops. The remarkable structural and functional organization of the eukaryotic pyruvate dehydrogenase complexes. Proc Natl Acad Sci USA 98:14802–14807, 2001. 27. DL Bates, MJ Danson, G Hale, EA Hooper, RN Perham. Self-assembly and catalytic activity of the pyruvate dehydrogenase multienzyme complex of Escherichia coli. Nature 268:313–316, 1977. 28. JH Collins, LJ Reed. Acyl group and electron pair relay system: a network of interacting lipoyl moieties in the pyruvate and alpha-ketoglutarate dehydrogenase complexes from Escherichia coli. Proc Natl Acad Sci USA 74:4223–4227, 1977. 29. LC Packman, CJ Stanley, RN Perham. Temperature dependence of intramolecular coupling of active sites in pyruvate dehydrogenase multienzyme complexes. Biochem J 213:331–338, 1983. 30. MJ Danson, AR Fersht, RN Perham. Rapid intramolecular coupling of active sites in the pyruvate dehydrogenase complex of Escherichia coli: mechanism for rate enhancement in a multimeric structure. Proc Natl Acad Sci USA 75:5386– 5390, 1978. 31. RL Cate, TE Roche, LC Davis. Rapid intersite transfer of acetyl groups and movement of pyruvate dehydrogenase component in the kidney pyruvate dehydrogenase complex. J Biol Chem 225:7556–7662, 1980. 32. DD Jones, KM Stott, MJ Howard, RN Perham. Restricted motion of the lipoyllysine swinging arm in the pyruvate dehydrogenase complex of Escherichia coli. Biochemistry 39:8448–8459, 2000. 33. HJ Chauhan, GJ Domingo, HI Jung, RN Perham. Sites of limited proteolysis in the pyruvate decarboxylase component of the pyruvate dehydrogenase multienzyme complex of Bacillus stearothermophilus and their role in catalysis. Eur J Biochem 267:7158–7169, 2000.

20 Activation and Transfer of Lipoic Acid in Protein Lipoylation in Mammals Kazuko Fujiwara, Kazuko Okamura-Ikeda, and Yutaro Motokawa University of Tokushima, Tokushima, Japan

I. INTRODUCTION Lipoic acid is a disulfide-containing cofactor widely distributed among living organisms (Fig. 1). It attaches to the acyltransferase subunit (E2) of the pyruvate, a-ketoglutarate, and branched-chain a-ketoacid dehydrogenase complexes and H-protein of the glycine-cleavage system via an amide linkage between the carboxyl group of lipoic acid and the q-amino group of a specific lysine residue of these proteins (1–4). As mentioned in the other chapters, the lipoyllysine arm on E2 and H-protein shuttles the reaction intermediate and reducing equivalents between the active sites of the complexes in the reaction sequence. Reed and coworkers first described the protein lipoylation in Streptococcus faecalis. The cell extracts from S. faecalis exhibited an activation of the apopyruvate dehydrogenase system prepared from mutant S. faecalis in the presence of lipoic acid, ATP, Mg2+, and inorganic phosphate. Because lipoylAMP could replace lipoic acid and ATP, the following two-step mechanism was proposed for the covalent attachment of lipoic acid (5): Lipoic acid þ ATP

!

lipoyl-AMP þ PPi

ð1Þ 343

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Figure 1 Structure of (R)-(+)-lipoic acid.

Lipoyl-AMP þ apoprotein

!

holoprotein þ AMP

ð2Þ

Lipoic acid is activated by Reaction (1), and then the lipoyl moiety is transferred to apoprotein by Reaction (2). Lipoate-protein ligase A purified from Escherichia coli and the recombinant enzyme catalyze both Reactions (1) and (2) (6,7). In contrast, lipoyltransferase purified from bovine liver catalyzes Reaction (2) but has no ability to activate lipoic acid to lipoyl-AMP (8,9). Thus, another enzyme, lipoate-activating enzyme, is required to activate lipoate in mammals (10). In this chapter, we describe the protein lipoylation mechanism in mammals. II. ACTIVATION OF LIPOIC ACID A. Purification and Characterization of Lipoate-Activating Enzyme The first study on mammalian protein lipoylation was carried out by Tsunoda and Yasunobu (11). Lipoate-activating enzyme was partially purified from the supernatant fraction of bovine liver homogenates by ammonium sulfate fractionation and calcium phosphate gel adsorption. The purified enzyme activated lipoic acid in the presence of ATP. Octanoic acid and lipoic acid derivertives were also activated by the enzyme. However, it had not been clarified whether the enzyme had the ability to transfer the activated lipoic acid to proteins because there was no appropriate apoprotein at that time. Recently, lipoate-activating enzyme (LAE) was purified from bovine liver mitochondria (10). Because Reaction (1) is closely similar to a partial reaction of acyl-CoA synthetase reactions, a coupled assay method was devised to isolate a specific enzyme involved in lipoate activation. The reaction mixture contained two enzymes, LAE and previously purified lipoyltransferase, in addition to lipoic acid, GTP, MgCl2, bovine apoH-protein, potassium phosphate buffer, pH 7.8, Tris-Cl buffer, pH 7.5, DTT, and bovine serum albumin. In this method, lipoate activated by LAE was immediately used for the lipoylation of apoH-protein by the action of lipoyltransferase. After the

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reaction, the amount of lipoylated H-protein was determined by the glycine–14CO2 exchange reaction (4). When the LAE activity was determined employing GTP as a high-energy compound, more than 99% of the LAE activity was confined to mitochondria (Table 1), whereas with ATP the activity was extremely low and distributed mainly in the microsomal fraction. Then LAE was purified from bovine liver mitochondrial extracts by chromatographies on DEAE-Sepharose, hydroxylapatite, blue-Sepharose, and Superdex 200 columns to apparent homogeneity, with a yield of 16% and a 39-fold purification. Throughout the purification steps, a single peak of LAE activity was detected by the coupled method, and elution profiles of the activity determined by a hydroxamate method (10,11) were parallel with those determined by the coupled method, suggesting that the purified LAE is primarily responsible for the activation of lipoic acid. Molecular masses of 61 kDa and 49 kDa were determined by SDS-PAGE and gel filtration chromatography, respectively. pI of 5.67 was determined by chromatofocusing, although theoretical pI was calculated to be 6.41 by the DNASIS program. Lipoic acid contains a chiral carbon at position C-6 (Fig. 1); therefore, (R)-(+)- and (S)-(-)-enantiomers are possible. (R)-lipoic acid is a naturally occurring enantiomer of lipoic acid. The LAE reaction was dependent on (R)lipoate, GTP, and MgCl2, because removal of (R)-lipoate, GTP, or MgCl2 from the reaction mixture resulted in no holoH-protein formation. With (S)lipoate, holoH-protein formation determined by the glycine–14CO2 exchange reaction was less than 1% of that with (R)-lipoate. Removal of lipoyltansferase resulted in no holoH-protein formation, indicating that LAE has no ability to transfer the activated lipoate to the apoprotein. By the coupled method, Vmax values with GTP, CTP, or UTP were about 1000-fold greater than that with ATP (Table 2). However, the activity with ATP determined by the hydroxamate method was about 100-fold higher than that determined by

Table 1 Subcellular Distribution of LAE Activity Determined by the Coupled Method Activity Fraction Mitochondria Microsome Cytosol

With ATP

With GTP

nmol/h/g liver 0.18 2.15 0.01

nmol/h/g liver 909.09 4.12 0

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Fujiwara et al. Table 2 Kinetic Constants for LAE Determined by the Coupled Method with Various Nucleotides Varied substrate ATP (R)-lipoate GTP (R)-lipoate CTP (R)-lipoate UTP (R)-lipoate

Km(app) mM 1.3 0.0047 0.37 0.041 1.2 0.039 8.8 0.18

Vmax(app) nmol/h/mg of protein 14.4 14.3 13800 15300 13500 12700 14100 13000

Source: Ref. 10.

the coupled method (data not shown). These results indicate that lipoyl-AMP is hardly released from LAE, while lipoyl-GMP, lipoyl-CMP, and lipoylUMP are easily released from LAE and served as substrates for lipoyltransferase. The Km value for GTP is lower than that for ATP. The concentration of GTP in mitochondria is reported to be in the range from 0.15 to 0.23 mM (12). These results strongly support a concept that GTP is involved in the activation of lipoic acid in mitochondria. The product of the LAE reaction was confirmed to be lipoyl-GMP by an HPLC analysis on an ODS-column (Fig. 2). The LAE reaction was carried out with (R)-lipoate or (S)-lipoate in the presence of GTP, MgCl2, DTT, TrisCl buffer, pH 7.5, and 0.249 Ag of LAE. After the reaction, the products were

Figure 2 Analyses of the reaction products. (A) HPLC analysis of LAE reaction products. The LAE reaction was carried out with (R)-lipoate (c) or (S)-lipoate (d) in the presence of LAE, or with (R)-lipoate in the absence of LAE (b). After reaction, the products were analyzed on an ODS-column employing an acetonitrile gradient with medium A (0.05 M sodium phosphate buffer, pH 5.5) and medium B (acetonitrile). Arrows indicate retention times of the standard lipoyl-GMP (a). —, absorbance at 252 nm; ---, concentration of acetonitrile. (B) Native-PAGE analysis of H-protein lipoylated with lipoyl-GMP obtained in panel A. ApoH-protein was lipoylated by lipoyltransferase without lipoyl-GMP (lane 1) or with (R)-lipoyl-GMP (lane 2) or (S)-lipoyl-GMP (lane 3) and analyzed on a native-PAGE. (C) The glycine–14CO2 exchange activity of the lipoylated H-protein. ApoH-protein (142 pmol) was lipoylated by 0.33 Ag of lipoyltransferase with 20 pmol of (R)-lipoyl-GMP (column 2) or (S)-lipoyl-GMP (column 3) obtained in panel A or without lipoylGMP (column 1). (From Ref. 10.)


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resolved on an ODS-column with an acetonitrile gradient. Authentic lipoylGMP elutes at 20.9, 23.9, and 25.6 min. The difference in the retention time presumably depends on the ionized states of lipoyl-GMP. The retention times of the reaction products with (R)- and (S)-lipoate were similar to those of the authentic lipoyl-GMP (Fig. 2A, c and d). The rates of (R)- and (S)-lipoylGMP formation were calculated to be 2475 and 1597 pmol/h, respectively. LAE showed higher affinity for (R)-lipoate than for (S)-lipoate (data not shown). Thus LAE activates both (R)- and (S)-lipoate but has a preference for (R)-lipoate. B. Lipoate-Activating Enzyme is a Mitochondrial Medium-Chain Acyl-CoA Synthetase cDNA clones for bovine LAE were isolated. The nucleotide sequence and the predicted amino acid sequence are shown in Figure 3 (DDBJ/EMBL/GenBank accession number AB048289). The cDNAs contain an open reading frame encoding a protein of 577 amino acids, including a mitochondrial presequence of 31 amino acids. A hypothetical ATP/AMP-binding motif typically found in an acyl-CoA synthetase family is shown. Amino acid sequence homology search by FASTA and BLAST showed that the sequence was identical to that of xenobiotic-metabolizing medium-chain fatty acid:CoA ligase-III from bovine liver reported by Vessey et al. (13,14). However, the isolated cDNA extends by 28 nucleotides upstream from the 5V-end of the reported cDNA and contains the following two differences from the reported sequence. First, a single nucleotide substitution of G for A due to SNP at position 1244 was found in four out of five clones analyzed, and consequently an amino acid substitution of alanine for threonine was predicted. Second, an insertion of 39 nucleotides due to an alternative splicing was found in two out of five clones. The purified LAE exhibited activities of medium-chain acyl-CoA synthatase, with the highest activity being with hexanoic acid (Fig. 4A). AcylGMP formation showed broad substrate specificity with respect to the chain length of fatty acid. The rate of lipoyl-GMP formation was comparable with that of decanoyl-GMP formation. Synthesis of hexanoyl-CoA was reduced by addition of lipoate and GTP (Fig. 4B). The inhibition by lipoate was competitive with hexanoate, and inhibition by GTP was competitive with ATP, indicating that GTP and ATP share a nucleotide-binding site and lipoate and hexanoate share a fatty acid–binding site on LAE. Similarly, lipoyl-GMP formation was inhibited by substrates for acyl-CoA synthesis (Fig. 4C). The addition of ATP inhibited the reaction nearly completely, because lipoyl-AMP formed hardly dissociated from LAE. However, the inhibition by ATP was slightly recovered in the presence of CoA. Although


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Figure 3 Nucleotide sequence of cDNA for LAE and the deduced amino acid sequence. The nucleotide numbering starts at the 5V end of the longest cDNA. The amino acid numbering starts at the initiation methionine. The amino acid sequence is shown in a single-letter code below the nucleotide sequence. The predicted mitochondrial presequence is underlined. Putative ATP/AMP binding motif is doubly underlined. An insertion of nucleotides (nucleotides 80–118) is shaded. A nucleotide change from A to G at position 1244 and a consequent amino acid substitution of T by A are indicated in black boxes. The asterisk represents the stop codon.

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Figure 4 Acyl-CoA synthesis and acyl-GMP formation catalyzed by LAE. (A) Substrate specificity with respect to chain length of fatty acid. ., less than 1% of the octanoyl-GMP formation; *, not determined. (B) Inhibition of hexanoyl-CoA synthesis by substrates of lipoyl-GMP formation. (C) Inhibition of lipoyl-GMP formation by substrates of hexanoyl-CoA synthesis.

Figure 5 Reaction catalyzed by LAE/medium-chain acyl-CoA synthetase. LAE and medium-chain acyl-CoA synthetase (MACS) are identical enzyme. In the acylCoA synthetase reaction, ATP is an essential substrate, whereas LAE utilizes GTP in the activation of lipoate, and the product is released from LAE to provide a substrate for lipoyltransferase. Then lipoyltransferase lipoylate apoproteins employing lipoylGMP.


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the rate of lipoyl-GMP formation in this situation is very low, it is still about 40-fold greater than that of lipoyl-AMP formation. These results suggest that in mitochondria where ATP, CoA, and GTP coexist, LAE can catalyze the lipoyl-GMP formation. However, the presence of some biochemical mechanism responsible for the preferential production of lipoyl-GMP may be required in mitochondria. The LAE reaction is summarized in Figure 5. Lipoate-activating enzyme is identical to the mitochondrial medium-chain acyl-CoA synthetase. In the acyl-CoA synthetase reaction, ATP is an essential substrate, because the reaction intermediate, acyl-AMP, has to be retained on the active site of the enzyme to react with CoA to produce acyl-CoA, whereas LAE catalyzes the activation of lipoate, utilizing GTP, and the product is easily released from the active site of the enzyme to provide a substrate for lipoyltransferase. Then lipoyltransferase lipoylates apoproteins using the lipoyl-GMP.

III. TRANSFER OF LIPOIC ACID TO PROTEINS A. Properties of Lipoyltransferase Our study on lipoyltransferase preceded the study on LAE. Therefore, the enzyme was purified from bovine liver mitochondria employing lipoyl-AMP and apoH-protein in an assay mixture as a donor and an acceptor of lipoic acid, respectively (4). The reaction mixture contained lipoyl-AMP, bovine apoH-protein, potassium phosphate buffer, pH 7.8, bovine serum albumin, and lipoyltransferase. After the reaction, the activity of lipoylated H-protein was determined by the glycine–14CO2 exchange reaction. The lipoyltransferase reaction was absolutely dependent on lipoyl-AMP, apoH-protein, and lipoyltransferase and was stimulated in the presence of bovine serum albumin and phosphate ion (8). Lipoyltransferase has no ability to activate lipoic acid to lipoyl-AMP because lipoic acid and ATP could not replace lipoyl-AMP. The purified enzyme utilized lipoyl-GMP as a donor of lipoic acid as well (10). (R)- and (S)-lipoyl-GMP eluted from the ODS-column was isolated separately (Fig. 2A, c and d), and 20 pmol of them was subjected to lipoyltransferase reaction employing 0.33 Ag of the enzyme and 7.1 pmol of apoH-protein. Then the lipoylated H-protein was resolved on native-PAGE (Fig. 2B). Lipoylated H-protein migrates faster on native-PAGE than apoHprotein because of the reduction of a positive charge of the lysine residue to be lipoylated. Lipoyltranferase equally transferred both (R)- and (S)-lipoyl moiety from respective lipoyl-GMP to apoH-protein. However, (S)-lipoylated H-protein did not exhibit the glycine–14CO2 exchange activity (Fig. 2C). Thus, it was clearly demonstrated that only H-protein carrying (R)-lipoyl moiety was active in the glycine-cleavage reaction.

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The purified lipoyltransferase also transferred lipoic acid to lipoyl domains of E2s of rat pyruvate, rat a-ketoglutarate, and bovine branchedchain a-ketoacid dehydrogenase complexes (abbreviations are PDC, KGDC, and BCKDC, respectively) in addition to bovine H-protein (15). However the lipoylation efficiency for the lipoyl domain of BCKDC was extremely low. Comparison of amino acid sequences surrounding the lipoic acid attachment site of the lipoate-acceptor proteins shows that the lipoyl domain of bovine BCKDC lacks the Glu residue at the three-residues N-terminal side of the lipoylation site (16) (Fig. 6). The Glu residue at the position is highly conserved among PDC (17–21), KGDC (22–24), and H-proteins (25–32), while Gln substitutes for the Glu residue in BCKDCs (16,33,34). Replacement of the Gln residue at the position of bovine BCKDC by Glu (Q41E) resulted in about 80-fold stimulation of lipoylation rate. On the other hand, replacement of the Glu residue at the position of rat PDC (17) and KGDC

Figure 6 Comparison of amino acid sequences surrounding the attachment site of lipoic acid. The sequences of dihydrolipoamide acetyltransferase of rat (17), human (18), Saccharomyces cerevisiae (19), E. coli (20), and Azotobacter vinelandii (21), of dihydrolipoamide succinyltransferase of rat (22), E. coli (23), and A. vinelandii (24), of acyltransferase of BCKDC of bovine (16), human (32), and chicken (33), and of H-protein of bovine (25), human (26,27), chicken (28), pea (29,30), and E. coli (31) are shown. The gray boxes show the lipoic acid attachment sites. The black boxes show the conserved amino acid residues interested. The numbers on the right and left refer to the position of the amino acid in the proteins.

Figure 12.1 Ribbon drawing of a complete YPDC subunit showing the triangular relationship between domains. The TDP, Mg2+, and C221 side chain are shown as space-filling representations. The coordinate set used was 1PVD. The figure was made with the program RIBBONS (Ref. 63).

Figure 12.2 Ribbon drawing showing the tightly associated YPDC dimer pair. The TDP and Mg2+ cofactors are included with a space-filling representation and lie at the interface between subunits. The coordinate set used was 1PVD. The figure was made with the program RIBBONS (Ref. 63).

Figure 12.3 Ribbon drawing of the complete YPDC tetramer in the ‘‘open’’ Form A viewed down the crystallographic twofold axis. The TDP and Mg2+ cofactors are included with a spacefilling representation. The center of the tetramer is open, and all dimer–dimer contacts are between the h domains that form extended 14-strand h sheets. The inset shows that at the interface the h strands are essentially perpendicular. The coordinate set used was 1PVD. The figure was made with the program RIBBONS (Ref. 63).

Figure 12.4 Ribbon drawing of the complete YPDC tetramer in the ‘‘closed’’ Form B. The TDP and Mg2+ cofactors are included with a space-filling representation. Those residues that become ordered in Form B as compared to Form A are shown in ball-and-stick representation. Note that in moving from Form A to Form B one side of the tetramer retains its h strand contacts but a new set of contacts is formed on the other side. The coordinate set used was 1QPB. The figure was made with the program RIBBONS (Ref. 63).

Figure 12.5 Least squares alignment of the structures of YPDC crystallized in the presence of ketomalonate (yellow) and pyruvamide (green). Note that although each structure was in a different space group, the structures are essentially identical. The figure was made with the program RIBBONS (Ref. 63).

Figure 12.6 Ball-and-stick representation of the active site of YPDC in Form A. Interactions shown are those involved in stabilizing the V-conformation or likely to be involved in catalysis. Residues numbered over 390 are from the g domain of one subunit; those under 115 are from the a domain of the ‘‘other’’ subunit. The figure was made with the program RIBBONS (Ref. 63).

Figure 12.8 Ribbon drawing of the activation pathway between the regulatory site and the active site. TDP and residues forming the information transfer pathway are shown in ball-and-stick representation. The figure was made with the program RIBBONS (Ref. 63).

Figure 12.9 Ribbon drawing of the alternate activation pathway, which could trigger the breakage of the interactions between the h sheets from loosely associated dimer pairs. One subunit ribbon is shown in yellow, the other in blue. Key residues are shown in ball-and-stick representation. T320 main-chain atoms form the hydrogen bonds at the dimer interface. The figure was made with the program RIBBONS (Ref. 63).

Figure 16.3 Model of the active site of BAL based on the sequence/structure alignments shown in Figure 2.

Figure 16.4 Overlay of active-site residues of BAL with those of (A) BFD and (B) ZmPDC.

Figure 23.1 (a) Ribbon drawings of the N-terminal, middle, and C-terminal domains (left to right) in the E. coli PHDc E1 subunit. (b) Ribbon drawings of the complete E1 subunit (left) and functional a2 homodimer (right). (c) Stereo drawing of the E1 active-site environment, including the TDP (in green). Residues numbered lower than 471 are from the N-terminal domain of one subunit, whereas those numbered greater than 470 are from the middle domain of the ‘‘other’’ subunit in the dimer. The main chain and H407 residue unobserved in E1 but positioned via least squares alignment with TK is shown colored in magenta. Several water molecules are included. Figures a and b were created with the program MOLSCRIPT (Ref. 27), Figure c was created with the program RIBBONS (Ref. 28).

Figure 23.2 (a) Stereo superposition of the E. coli PDHc E1 subunit with yeast transketolase after least-squares alignment. Colors are black and green for the E1 and TK structures, respectively. Two TDP molecules are shown in blue. (b) Stereo superposition of the E. coli PDHc E1 subunit with its P. putida E1 counterparts, after least-squares alignment. Colors are as in part (a). Both figures a and b were created with the program MOLSCRIPT (Ref. 27).

Figure 23.4 Location of tyrosine 177 and histidine 179 with respect to TDP in the E. coli PDHc E1 active site. Inset: Progress curve for NADH production in the overall PDHc reaction in the presence of H179A PDHc-E1 and 0.10 mM or 0.40 mM TDP.


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(22) by Gln (E169Q and E40Q, respectively) resulted in a reduction of lipoylation of less than 0.3% of the lipoylation rate of the wild types. Three-dimensional structures of the lipoyl domains of PDC (35–37) and KGDC (38,39) from various species and pea H-protein (40) have been resolved. They form a similar h-barrel structure consisting of two almostparallel four-stranded antiparallel h-sheets formed around a well-defined hydrophobic core. The lipoylation site is exposed to the surface of the domain in a tight h-turn in one of the antiparallel h-sheets. The Glu residue is also exposed to the surface, just at the end of the preceding h-sheet. These results indicate that the Glu residue at the position plays an important roll in recognition of the lipoylation site by lipoyltransferase. Wallis and Perham reported a requirement of accurate positioning of the target Lys residue in the h-turn for the recognition by E. coli lipoate-protein ligase A (41). The positional relationship between the Lys and Glu residues may be important for the recognition of the lipoylation site by lipoyltransferase and lipoateprotein ligase A. Replacement of the Gly residue by Ser or Asn at the 11-residues Cterminal side of the lipoylation site of rat KGDC (G54S or G54N) resulted in a reduction of the mobility of the lipoyl domain on a native-PAGE. Moreover, G54N showed no lipoylation (15). Because the Gly residue is located at the opposite side of the lipoylation site in the lipoyl domain, the residue seems to be important to keep a proper conformation required for the lipoylation. Although the Gly residue at the 16-residues N-terminal side of the lipoylation site is well conserved, replacement of the residue by Ser or Asn did not show any effect on the mobility on the native-PAGE or the lipoylation of the domain. Therefore, the Gly residue seems not to be involved in the recognition by lipoyltransferase. B. cDNA and Genomic DNA Structures of Lipoyltransferase cDNAs for bovine and human lipoyltransferase were cloned (9,42). The nucleotide sequence data are available (DDBJ/EMBL/GenBank database with accession numbers AB006441 and AB017566, respectively). The predicted amino acid sequences and the sequence of E. coli lipoate-protein ligase A are aligned in Figure 7. Bovine and human lipoyltransferases contain a mitochondrial targeting sequence of 26 amino acids. They share 88% identity in the amino acid sequences. The amino acid sequence of E. coli lipoate-protein ligase A shares 31% and 33% identity with those of human and bovine enzymes, respectively. In particular, amino acids 34–112 of mammalian lipoyltransferases share high homology with amino acids 6–85 of the E. coli enzyme. It suggests that the N-terminal half of these proteins may be responsible for the lipoate-transferring activity and the C-terminal half of the

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Figure 7 Sequence alignment of lipoyltransferases (LT) (9,42) and E. coli lipoateprotein ligase A (Lp1A) (6). Amino acid numbering indicated on the right starts at the initiation methionine. The asterisk represents an N-terminus of the mature lipoyltransferases. Amino acid residues identical to the human enzyme are indicated in black boxes.


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E. coli enzyme may contribute to the lipoate-activating activity that mammalian enzymes lack. Genomic DNA for the human lipoyltransferase was cloned (42). The nucleotide sequence data can be found in DDBJ/EMBL/GenBank database with accession number AB017567. The gene is organized into four exons and three introns spanning about 8 kbp of genomic DNA (Fig. 8). Exon IV contains the entire coding sequence. Multiple alternative splicings were found in the 5V-noncoding region by 5V-RACE analysis. Transcript A was the most abundant species. Significance of the alternative splicing, such as tissue dependence or relationship with disease, has not been clarified. Mapping of

Figure 8 Restriction map and exon-intron organization of the human lipoyltransferase gene. (A) The structure of the lipoyltransferase gene. Exons are indicated by rectangles and the coding region is indicated by a closed rectangle. SacI (S), PstI (P), HindIII (H), and EcoRI (E) restriction sites are shown. (B) Schematic diagrams showing the heterogeneity of the 5V-untranslated region of the transcripts (designated transcripts A–D) defined by 5V-RACE analysis. (From Ref. 42.)

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the human lipoyltransferase gene by fluorescence in situ hybridization showed that the gene was localized to the q11.2 band of chromosome 2 (42). The expression of mRNA of lipoyltransferase in various human tissues was examined by Northern blot analysis (Fig. 9A). The transcript was about 1.5 kb in size and expressed in all tissues examined. However, the expression was highly regulated and most abundant in skeletal muscle and heart. Interestingly, similar patterns of expression of the lipoate acceptor proteins, E2 of PDC, KGDC, and BCKDC and H-protein, were observed (Fig. 9B–E). They were also highly expressed in skeletal muscle and heart, as reported previously (18,43). This correlation enables lipoyltransferase to respond to the requirement of lipoylation of these proteins.

Figure 9 Northern blot analyses of the human lipoyltransferase mRNA and mRNAs of lipoate acceptor proteins. Northern blot from human multiple tissue was analyzed with probes corresponding to the cDNAs for the human lipoyltransferase (A), E2 of the rat PDC (B) (17), E2 of the rat KGDC (C) (22), E2 of the bovine BCKDC (D) (16), and bovine H-protein (E) (25). The blot was also hybridized with a human h-actin cDNA as a control to ascertain the difference in RNA loaded (F). Molecular mass markers are indicated in kilobases on the left. Pa, pancreas; K, kidney; S, skeletal muscle; Li, liver; Lu, lung; Pl, placenta; B, brain; H, heart. (From Ref. 42.)


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IV. SUMMARY In mammals, lipoic acid supplied via the diet is incorporated into mitochondria through some transporters (44–46). In mitochondria, lipoic acid is activated to lipoyl-GMP by LAE, employing GTP as a high-energy compound; then the lipoyl moiety is transferred to apoproteins by the action of lipoyltransferase (Fig. 10). On the other hand, E. coli lipoate-protein ligase A activates lipoic acid, employing ATP, retains the intermediate, lipoyl-AMP, and transfers the lipoyl moiety to apoproteins (6,7). Thus, lipoate-protein ligase A catalyzes both the activation and the transfer reactions. In E. coli, another lipoylation pathway has been established. In this pathway, lipoic acid synthesized by the action of Lip A (47,48) from octanoic acid on the acyl carrier protein is transferred to apoproteins by Lip B (48,49), whereas lipoateprotein ligase A functions for incorporated exogenous lipoic acid. Recently,

Figure 10 Metabolism of lipoic acid in mammalian mitochondria and E. coli. (A) Lipoic acid incorporated into mitochondria is activated to lipoyl-GMP by LAE, and then the lipoyl moiety is transferred to apoproteins by lipoyltransferase (LT). (B) Lipoic acid incorporated into E. coli is activated with ATP and transferred to proteins by the action of lipoate-protein ligase A (Lp1A). Another protein lipoylation pathway is presented in E. coli, in which LipB transfers lipoyl moiety from lipoylacyl carrier protein (lipoyl-ACP) synthesized endogenously from octanoyl-ACP by LipA.

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21 Central Organization of Mammalian Pyruvate Dehydrogenase (PD) Complex and Lipoyl Domain–Mediated Activated Function and Control of PD Kinases and Phosphatase 1 Thomas E. Roche, Yasuaki Hiromasa, Ali Turkan, Xiaoming Gong, Tao Peng, Xiaohua Yan, Shane A. Kasten, Haiying Bao, and Jianchun Dong Kansas State University, Manhattan, Kansas, U.S.A.

I. INTRODUCTION A. Composition and Roles The mitochondrial pyruvate dehydrogenase complex (PDC) catalyzes the irreversible conversion of pyruvate to acetyl-CoA along with the reduction of NAD+. The components that are required for the overall reaction include the pyruvate dehydrogenase (E1) component, the dihydrolipoyl acetyltransferase (E2), the dihydrolipoyl dehydrogenase (E3) component, and the E3-binding protein (E3BP). Mammalian PDC has a highly organized structure in which the E2 component plays a central role in the organization, integrated chemical reactions, and regulation of the complex (1–4). The E3BP component also contributes to these roles. This chapter presents new insights into the 363

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integration of E3BP into the central framework of the complex. The major focus is the predominate consequences for the regulation of PDC of the interplay between E2-E3BP and the kinase/phosphatase enzymes (see later) that govern PDC activity. PDC plays strategic fuel-selection roles in determining whether, under different metabolic conditions, glucose-linked substrates are converted to acetyl-CoA in mammalian tissues. PDC activity is up-regulated for the use of glucose-linked substrates as sources of oxidative energy or as precursors in the biosynthesis of fatty acids (3–9). Intake of excess dietary carbohydrate leads to insulin-mediated activation of PDC activity in fat-synthesizing tissues to promote the biosynthesis of fatty acids from glucose. The PDC reaction is a critical step in the aerobic use of glycogen stores and blood glucose by muscle tissues and is required for the standard glucose consumption in neural tissues. When carbohydrate stores are reduced, mammalian PDC activity is downregulated to limit the oxidative utilization of glucose in most nonneural tissues. Extended starvation causes PDC activity to be emphatically suppressed in most tissues. The same regulatory control severely confines PDC activity in diabetic animals to thereby obstruct consumption of abundant glucose. Adaptable control of PDC activity is required to satisfy these discrete roles in the management of fuel consumption and storage. B. Control and General Properties of Regulatory Enzymes The foregoing fuel-management role is achieved by tissue-specific and metabolic state–specific expression and the discrete regulatory properties of the dedicated protein kinase and protein phosphatase isozymes (1–12). Four pyruvate dehydrogenase kinase (PDK) isozymes and two pyruvate dehydrogenase phosphatase (PDP) isoforms function in governing the activity state of PDC (10–12). In combination these carry out a continuous phosphorylation– dephosphorylation cycle that determines the proportion of the pyruvate dehydrogenase (E1) component that is in the active, nonphosphorylated state. PDK isozymes, together with the related branched-chain dehydrogenase kinase, comprise a novel family of serine kinases, unrelated to cytoplasmic Ser/Thr/Tyr kinases (3,4,6–11,13–16). Based on the order in which they were initially cloned, the four PDK isoforms identified in mammals are designated PDK1, PDK2, PDK3, and PDK4 (10,11). The PDKs have two-domain structures; the C-terminal domain is clearly related to another class of ATP-consuming enzymes (13–16) that broadly includes bacterial histidine kinases. The sequences of the same isozyme in different mammals are highly conserved (>94% identity for human versus rat) (3,11,13,14). The different 45.5- to 46-kDa human isoforms share 65% F 4% sequence identity with only short segments at the N-terminus that cannot be aligned.

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To conserve carbohydrate reserves, feedback suppression of the PDC reaction when fatty acids and ketone bodies are being used as preferred energy sources results from enhanced kinase activity (1–5). The resulting elevation of the ratios of intramitochondrial NADH to NAD+ and acetyl-CoA to CoA suppresses PDC activity by effectively stimulating kinase activity. Kinase activity is reduced by ADP, which when elevated evinces a lowenergy state, and by pyruvate, which manifests the availability of substrate. Kinase activity is synergistically inhibited by a combination of these effectors (17). Inorganic phosphate anion also fosters ADP and pyruvate inhibition. As described elsewhere in this volume and in previous studies (7,8,18– 22), Harris, Sugden, and coworkers have characterized the control of the expression of the PDKs. Particularly important is the overexpression of PDK4 during starvation, which spawns a need to conserve carbohydrate reserves. PDK4 expression is increased both by glucocorticoids and by free fatty acids via the peroxisome proliferator–activated receptor a and is hindered by an insulin-activated pathway (7,21). In diabetic animals, the unimpeded functioning of the latter regulatory pathway (due to lack of insulin or insensitivity to insulin) deleteriously fosters overexpression of PDK4 to thereby curtail glucose oxidation. The two PDP isoforms have 52-kDa catalytic subunits that are members of the 2C class of protein phosphatase (12,23). Besides its catalytic subunit (PDP1c), PDP1 also contains a large (95.6-kDa) regulatory subunit (PDPr) that retains an FAD (24); PDPr shares a 35% sequence identity with the mitochondrial flavoprotein dimethylglycine dehydrogenase (25). Both PDP1 and PDP2 activities require Mg2+ and are regulated with regard to their responsiveness to this essential metal (12,23–29). Micromolar Ca2+ greatly stimulates the activity of PDP1, which is found in Ca2+-sensitive tissues (9,28). Polyamines, most especially spermine, significantly reduce the Km values for Mg2+ of both PDP isoforms (12,26). The Km of PDP2 in the absence of polyamines is very high (16 mM) and is reduced to 3 mM by spermine (12), whereas the Km of PDP1 for Mg2+ is lowered from 2 mM (+Ca2+) to 0.4 mM by spermine (26–29). PDP1r increases the Km of PDP1 for Mg2+ (26,29); PDP1c alone has a low Km for Mg2+, similar to sperminetreated PDP1 (23,29). It remains uncertain whether spermine levels change in mitochondria or whether polyamines mimic another intramitochondrial effector. PDP2 is expressed in fat-synthesizing tissues (12) and is probably the primary target by which insulin-predicated regulation enhances PDP activity via a mechanism that, like spermine, leads to a lowering of the Km for Mg2+ (30). Putative final-stage mechanisms whereby insulin regulation enhances PDP activity include allosteric mediators (31) and phoshorylation by PKCy (32).

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C. E2- and E3BP-Domain Structures and Domain Roles E2 and the E3BP component have remarkable structures that allow them to carry out their related general roles in forming the anchoring assemblage and integrating the sequential reactions of the complex. Mammalian PDC-E2 has four globular domains (Fig. 1), with two linker-region-connected lipoyl domains at its N-terminal end and an assembly-forming, catalytic domain at the C-terminal end (1–4,33,34). The N-terminal lipoyl domain is designated L1 and the second lipoyl domain L2. Between the C- and N-terminal domains is a small globular domain, flanked by linker regions, which binds the E1 component (35,36). When expressed by itself, 60 C-terminal domains associate as 20 trimers in the form of a pentagonal dodecahedron. In this structure, 12 pentagonal faces are created, with these 20 trimer corners associating to create 30 edges, each shared by two pentagonal faces (see Fig. 2 for the general form, although this structure is modified as described

Figure 1 E2 and E3BP domains and their binding interactions. The linker-regionconnected domains of the E2 subunit are: L1, the N-terminal lipoyl domain; L2, the inner lipoyl domain; B, the E1 binding domain; and I, the oligomer forming-, acetyltransferase-catalyzing inner domain. With related structures (see text) and connecting linker regions, the E3BP domains are: L3, the N-terminal lipoyl domain; B, the E3-binding domain; and I, the inner domain, which associates with the inner domain of E2. Dashed connections designate specific binding interactions of E2 and E3BP domains with other components (see text). Stronger binding is indicated by thicker dashes.


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later). Trimers act together to catalyze the transacetylation reaction in an active site formed between subunits (37,38). The E3BP component has a similar segmented structure, with globular domains connected by linker regions (1–4,34). E3BP associates with the inner E2 core via its C-terminal domain (39), binds E3 via a binding domain (34,35,40–43), and has a single Nterminal lipoyl domain (44,45) that is designated L3. Recent insights into the nature of this association of E3BP with E2 in the mammalian complex are described later. In all a-keto acid dehydrogenase complexes, lipoyl domains are mobile elements acting at the surface of the complex to consolidate the sequential five-step reaction sequence by serving as substrates in the three central reactions and as mobile carriers of the resulting intermediate forms of the lipoyl prosthetic group (oxidized disulfide, 6,8-dithiol, and 8-acetyl). The facile ability of the lipoyl domains to traverse between the E1, E2, and E3 active sites is furnished through the high mobility of the extended but somewhat stiff Ala-Pro–rich linker regions (46). A major focus of this chapter is the roles of E2 and to a lesser degree E3BP in carrying out specialized interactions within the PDC assemblage that enhance PDK and PDP1 activities and directly contribute to the selective processing of specific regulatory effects (1–4,24,36,47–58). Of central importance is the functional interplay of PDKs and PDP1 with the mobile lipoyl domains of the E2 component (Fig. 1). The lipoyl domain of the E3BP component contributes in this case one PDK. E2 transforms kinase and phosphatase function and regulation through functioning as an anchoring scaffold, an adaptor protein directly abetting efficient phosphorylation and dephosphorylation, a processing unit in translating and transmitting effector signals, and in altering the sensitivity to allosteric effectors, required cofactors, and reactants that bind directly to kinases and PDP1 (reviewed in Refs. 1–4). This chapter will emphasize pivotal mechanisms whereby E2 elicits the predominant changes in the operation and the effector modulation of the PDKs (emphasis PDK2 and PDK3) and PDP1. II. NEW INSIGHTS INTO THE ORGANIZATION OF MAMMALIAN PDC As already indicated, the C-terminal domain of E3BP was shown to associate with the inner core formed by E2’s C-terminal domain. The yeast E3BP could be added to the E2 60mer (59) by its C-terminal domain locating inside the dodecahedron (60). In contrast, resolved E3BP of the mammalian complex, which retained a capacity to bind E3 and had a functional lipoyl domain, failed to bind back to assembled E2 in the absence of chaotropic conditions (40,61,62). In the yeast complex, the C-terminal domains of the E2 and E3BP components were not related, based on their amino acid sequences (45). In

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marked contrast, the C-terminal domains of mammalian E2 and E3BP are closely related in sequence and size (34). The E3BP C-terminal domain lacks some residues that are essential for carrying out catalysis in the acetyltransferase reaction. Using recombinantly expressed human components in analytical ultracentrifugation (AUC) studies, we have found that the mass of E2 60mer is slightly larger than the mass of the E2-E3BP complex (Y. Hiromasa and T.E. Roche, unpublished). The E3BP subunit has a smaller mass than the E2 subunit (48,040 versus 59,551). Therefore, the lower mass of E2-E3BP seemingly suggests that the closely related C-terminal domains of E3BP substitute in the dodecahedron rather than adding to it. A striking AUC finding in support of this possibility was that E2 binds nearly 60 E1 tetramers, whereas the E2-E3BP binds about a dozen fewer E1 tetramers. Based on the estimated Mr of E3-saturated E2-E3BP-E3 complex, we find that E2-E3BP maximumly binds 12 E3 dimers. This favors the conclusion that there are about 12 E3BP in the complex. Numbers in this range have been previously suggested based on the addition to 60mer and the smallest symmetry element of 12 open pentagonal faces in the dodecahedron (60,63). Furthermore, the yeast E3BP apparently binds the E3 dimer in these open faces (60). We have also normalized all our protein measurements by simultaneous interference and 280-nm measurements in AUC studies and found standard procedures for measuring E2 protein tend to give 20% higher than this updated estimate. The procedures used previously were less quantitative and also measured component ratios in conjunction with assuming that there were 60 E2 subunits per core. If, as we estimate, the saturated E2-E3BP-E3 is 26% E3 by mass but we calculate that fewer complexes are present using the higher mass of the 60/12 model while taking into account the foregoing error in protein estimates, we calculate 16.8 E3 per core. Since the subunit composition E260E3BP12E324 has only 15.4% higher mass than E248E3BP12E324, this is within experimental error of the higher values reported previously. Often complexes are deficient in E3; only 70% of the level of E3 needs to be bound to estimate 12 E3 dimers per core with the preceding assumptions. Studies using small-angle X-ray scattering (SAXS)(Y. Hiromasa, T. Fujisawa, Y. Aso, T.E. Roche, unpublished) and cryoelectron microscopy (H.Zhou, J. Stoops, Y. Hiromasa, and T.E. Roche, unpublished) have not detected additional mass in the E2-E3BP structure that extends beyond the E2 60mer. Indeed, SAXS estimates, like AUC results, indicate that there is a smaller total mass for E2-E3BP than for E2. Therefore, the combination of a lower mass of E2-E3BP, the capacity to bind less E1, and the lack of mass exterior to the dodecahedron strongly supports a model in which E3BP subunits substitute for E2 subunits within the dodecahedron. That 12 E3 dimers are bound by E2-E3BP provides the best estimate of the level of E3BP present in the structure.


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We have evaluated potential models for an integrated E2-E3BP core with an objective of substituting E3BP C-terminal domain within the dodecahedron in a symmetry manner that matches the nearly constant level of E3BP that we have found over the years in preparations of bovine complex, bovine E2-E3BP subcomplex, and the recombinant human preparations. We have concluded that there is only one model that allows symmetric substitution while simultaneously fitting 20 or less E3BP being incorporated. We would first note that one E3BP cannot be added per trimer at all 20 corners of the dodecahedron to give symmetric positioning of E3BP. The minimal number of full trimer replacements (E3BP-trimers) that can be symmetrically introduced is eight, which would require 24 E3BP subunits. The model shown in Figure 2 introduces 12 E3BP in equivalent and symmetric positions. The first directive for assembly to fit this model is that

Figure 2 Proposed model of the inner core formed by E2 and E3BP. The inner domains (see Fig. 1) of E2 and E3BP are proposed to form a pentagonal dodecahedron with symmetrically distributed I domains of E3BP that form six-dimer connections (light balls). E2 inner domains (darker balls) connect only to E2 inner domains along the 2-fold axes. This view is tipped slightly off the twofold axis. In this view, one dimer representing E3BP inner domains is at the back of the domain and if the view was directly down, the twofold axis would be directly behind the front central E3BP dimer and four other dimers form edges at the sides of the structure. Four of the eight trimers formed only by E2 inner domains are modeled at the front of the structure; an equivalent set is at the back of the structure. The subunits of these trimers connect along the twofold axis with E2 subunits of the 12 trimers that contain one E3BP inner domain. See the text for further explanation of the organization.

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the C-terminal domains of E3BP self-associate as dimers that therefore connect two E3BP-containing trimers. This fits constraints being introduced by the distinct character of the parts of the domain that associate along the twofold axis. Recent electron microscopy results point to flexible ‘‘ball-andsocket’’ connections along the twofold axes between E2 subunits (64). Based on sequence alignments and the three-dimensional structures of the innercore dodecahedron of bacterial E2 (65), there are changes in the E3BP sequence in regions that are vital for forming this flexible interaction between E2 subunits. This includes the critical C-terminal residue (Leu or Met in E2’s) which forms the ‘‘ball’’ in a hydrophobic ‘‘ball-and-socket pocket’’ that allows flexible movement (extension and contraction) in the E2 connections along the twofold axis. Also, E3BP has a deletion of three residues in a region of H2 helix that contributes a part of the hydrophobic ‘‘socket pocket.’’ Thus, it seems unlikely that E3BP could form an interaction with E2 subunits along the twofold axis. However, a distinct but less flexible interaction with itself is likely, since there is still substantial sequence similarity of E3BP to E2 in the regions of E2 that associate along this axis. The second requirement is that the C-terminal domains of E2 in a trimer that contains an E3BP only associate with trimers that contain just E2. The preceding ‘‘ball-and-socket’’ connections along the twofold axis between E2 subunits expand and contract to cause substantial variation (up to 40 A˚) in the diameter of the inner core of the E2-oligomer. It has been proposed that this breathing contributes to the function of the complex. An additional possibility is that such flexibility might be required in the case of a substituted mammalian E248E3BP12 structure to compensate for differences in the length of the E3BP dimer connections along the twofold axis. The resulting model has 12 trimers containing one E3BP that associate to form six E3BP dimer connections and eight trimers containing only E2 that only associate via E2– E2 connections along the twofold axis with the 24 E2 subunits of the 12 E3BPcontaining trimers. Not only is this a viable model, but it is the only model that fits the requirement for symmetry and happens to simultaneously conform to the finding of a nearly constant level of 12 E3BP per complex. Further studies are under way to evaluate this model.

III. E2-FACILITATED KINASE FUNCTION A. Binding and Activation of PDK2 PDK2 is highly sensitive to the full set of known regulatory effects on mammalian PDK activities summarized earlier. The E2 component markedly increases the efficiency of PDK2 catalysis and intervenes to produce or modify all of these regulatory responses (56) (X. Yan, H. Bao, S.A. Kasten, and T.E.


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Roche, unpublished work). We will first consider how E2 supports greatly enhanced PDK2 catalysis. Functional studies indicated PDK2 preferentially interacts with the L2 domain of E2 (56). With micromolar levels of E1, E2 enhances the rate of phosphorylation of E1 by severalfold; with nM complexes, the estimated rate enhancement increases to 5000-fold for the complexes as compared to an equivalent amount of free E1 undergoing phosphorylation by the free PDK2 (Y. Hiromasa and T.E. Roche, unpublished work). Most assuredly, this results from PDK2’s gaining efficient access to many E2-bound E1 via the agile intervention of the outer domains of the E2 60mer greatly increasing kinase access to its bound substrate. A variety of functional studies and binding studies conducted using AUC support PDK2 preferentially interacting with the inner lipoyl domain (L2 domain, Fig. 1) of E2 via an interaction that requires the lipoyl prosthetic group (52,54–57; Y. Hiromasa and T.E. Roche, J Biol Chem, in press). In AUC studies, binding to a free L2 monomer is very weak (Kd f 80 lM), but binding to two L2 in GST-L2 dimer structure is appreciably tighter (Kd = 3.5 lM) (Y. Hiromasa and T.E. Roche, unpublished work). Therefore bifunctional binding of PDK2 dimer to two L2 is strongly supported. Our AUC studies firmly supported a much weaker binding of monomeric L2 to PDK2 than was previously estimated by a gel filtration approach (66). The strength of binding is increased more than 30-fold upon reduction of the prosthetic group (i.e., by GST-L2red). This finding has relevance for PDK2 regulation since lipoate reduction (and reductive acetylation) stimulates PDK2 activity (mechanism discussed later). There was much weaker binding to the reduced forms of GST-L1 and GST-L3 (see Fig. 1). Thus, these studies with free lipoyl domain structures predicts that PDK2 dimer binds E2 by associating with two lipoyl domains and that the affinity of binding will be increased when E2 lipoyl groups are reduced. Direct binding of PDK2 to E2 occurs at about 20 sites per E2 60mer, with an affinity similar to the binding to GST-L2 (Y. Hiromasa and T.E. Roche, J Biol Chem, in press). This further supports the importance of bifunctional binding. The presence of E2-bound E1 further strengthens the affinity of PDK2 for the complex by about 10-fold (Kd f 0.3 lM); this is consistent with the expected further interaction of PDK2 with its E1 substrate. However, this binding affinity is not sufficiently tight to fully explain how PDK2 activity is maintained at its maximum with dilute complexes (30 nM) containing less than 0.5 PDK2 dimers per complex (Kd = 0.3 lM and 20 sites/60mer predicts 62% of 0.5 PDK2 dimer per core bound at 30 nM). Inclusion of ADP or the ATP analog, AMP-PNP, an effective inhibitor of PDK2, and phosphorylation of E1 decreased binding of PDK2 to E2-E1 in AUC studies. Thus, kinetic features of the PDK2 reaction or other supporting mechanisms are needed to augment binding

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in order to explain how maximal PDK2 activity is maintained with 30 nM complex. B. Binding and Activation of PDK3 The free L2 domain, with an oxidized lipoyl group, binds PDK3 with a much tighter affinity than its binding to PDK2, and just the L2 domain strongly enhances PDK3 phosphorylation of free E1 (56). At 100-fold lower levels, the GST-L2 dimer similarly accelerates PDK3 activity (P. Tao, Y. Hiromasa, and T.E. Roche, unpublished work). PDK3 tends to form aggregates and the binding of two L2 stabilizes PDK3 as a dimer. Indeed, a portion of the 13-fold activation of PDK3 by L2 (56) results from preventing or reversing PDK3 self-association in the absence of L2. To obtain maximal activity, the standard procedure is to incubate dilute PDK3 with the E2-E1 complex for an extended period prior to the initiation of activity. This preincubation facilitates a 20- to 40-fold gain in PDK3 activity. Following 100-fold dilution of the L2stabilized PDK3 dimer into preinitiation assay mixtures using free E1, PDK3 maintains as high an activity as the nonaggregating PDK2. Further inclusion of L2 or GST-L2 still promulgates a several-fold higher kinase activity, suggesting that L2-binding induces a more active PDK3 conformation. Beyond this direct allosteric activation, the E2 60mer promotes higher PDK3 activity, and, in contrast to the enhancement by L2, high activity is maintained with very dilute complexes (1 mM (67,68), so locally several lipoyl domains can compete in the intramolecular step to allow rebinding to two lipoyl domains to occur faster than complete dissociation. Such a mechanism of facilitated access to E1 may be singularly important for sustained kinase function within the mitochondrion, where the high protein concentration (>400 mg/mL) limits diffusion of macromolecules. We have mapped the required surface of the L2 domain that is needed to bind and leverage a conformational change that elicits the large increase in PDK3 activity (X. Gong, T. Peng, and T.E. Roche, unpublished work). L2


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structures were produced that were modified both by substituting surface amino acid residues and by enzymatically adding cofactor analogs to the Lys that undergoes lipoylation. As shown in Figure 3A and B, a set of critical residues (kk-labeled for potent effects) were located near the lipoylated end of L2 (Leu140, Asp172, and Ala174, Asp197 and Arg196). Additional residues needed for activation were the acidic residues Glu162 and Glu179 (Fig. 3A), which are located more toward the other end of the domain. Even very high levels of the well-folded and fully lipoylated E179A-L2 failed to activate PDK3, suggesting that Glu179 is particularly important for leveraging the change in conformation that yields this activation. Substitutions of several different amino acids for the lipoylated Lys173 failed to activate PDK3. 8Thiol-octanoyl-L2 enhanced PDK3 activity beyond the native lipoylated lipoyl-L2. Heptanoyl-Lys173-L2 inhibited PDK3 activity and effectively hindered activation by native L2. Thus, it would appear that the full length of the lipoyl-lysine prosthetic group was absolutely required for leveraging PDK3 activation and that critical residues are widely distributed on the surface of the L2 domain. Therefore, there are ample interactions for both binding and altering the conformation of PDK3. The extended reach of the 8thiol group upon lipoate reduction is proposed to contribute to additional interactions that foster higher kinase activity. This is consistent with this mechanism’s producing kinase stimulation by NADH (see next section). C. E2-Mediated and -Modified Regulation of Kinase Activity As indicated in Section I, the consumption of fatty acids and ketone bodies as the primary energy source results in feedback suppression of PDC activity. This arises from kinase activity’s being appreciably enhanced due to the elevation of NADH/NAD+ and acetyl-CoA/CoA ratios as these fuels are consumed. The increase in these ratios is sensed and translated by the rapid and reversible E3 and E2 reactions, which increase the proportion of the lipoyl groups of E2 and E3BP that are reduced and acetylated (Fig. 4) (49,55– 57,69–74). Short-term reduction of the lipoyl group spawns up to an 80% increase in kinase activity (PDK*, Fig. 4); longer periods of reduction prior to initiating kinase activity apparently give higher levels of stimulation (73). Acetylation by further use of acetyl-CoA in the E2 reaction stimulates kinase activity up to threefold (PDK**, Fig. 4). In the absence of CoA, full stimulation can also be achieved via low levels of pyruvate reacting through the rate-limiting E1 reaction (49,55,70). Indeed, this means provided important mechanistic insights into the basis for stimulation, since blocking E1 catalysis prevented reductive acetylation and consequently kinase activation. Stimulation still takes place with peptide substrates and, in the absence of E2, with free lipoyl domains (55). This establishes the importance of direct

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Figure 4 Mechanism by which elevation of the ratios of NADH to NAD+ and acetyl-CoA to CoA lead to stimulation of PDK activity. In this condensed presentation, the E1, E2, and E3 interconvert lipoyl groups between the oxidized, reduced, or acetylated forms. The E1 reaction is rate limiting under most conditions, and the relative proportion of reaction states of the lipoyl groups is determined primarily by the rapid and reversible E3 and E2 reactions. Subsequent to those reactions, when a kinase dimer binds to a lipoyl domain with its lipoyl group in the oxidized form (bound PDK, top left), E2 facilitates higher rates of phosphorylation of the E2-bound E1. Binding of a kinase subunit to a lipoyl domain containing a reduced lipoyl group fosters a further increase in kinase activity (PDK* state). Finally, association of a kinase subunit with a lipoyl domain with a reductively acetylated lipoyl group leads to a further enhancement of kinase activity (PDK** state). See the text for the magnitude of these effects.

allosteric interactions of the reacted lipoyl group with the kinase. The inner E2 core stripped of lipoyl domains catalyzes the acetylation of 8-thiol-octanoylL2. This acetylation acts to stimulate kinase activity (T. Peng and T.E. Roche, unpublished). Therefore, the thiol at the 6-position of the dihydrolipoyl group either in just the reduced or in the acetylated form is not required for stimulation.

Figure 3 Surface residues of the L2 domain of E2 that are required for activating PDK3 and binding PDP1. Panels A and B present opposite sides of space-filled models of the human L2 domain. Lys173, which undergoes lipoylation, is located at the top of the structures; Lys-173 must be lipoylated for an L2 construct to enhance PDK3 activity or to bind and competitively prevent E2 activation of PDP1 (or PDP1c). In panels A and B, the supercript kk or k designates residues for which their substitution eliminates z80% or z50%, respectively, of the capacity of L2 to activate PDK3 (X. Gong, T. Peng, and T.E. Roche, unpublished work). Similarly, the superscript pp or indicates a site of substitution that reduces PDP1 binding to L2 by z75% or z45%, respectively (Ref. 76).

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Half-maximal stimulation of the activity of bovine kidney medulla PDK is achieved when a small proportion of the lipoyl groups (

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