Thiamine: Catalytic Mechanisms in Normal and Disease States (Oxidative Stress and Disease)

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

xvi

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

xxiii

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

2

Frey

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.

4

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-

Chemical Intermediates in Catalysis by TDP

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

Chemical Intermediates in Catalysis by TDP

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

Chemical Intermediates in Catalysis by TDP

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-

Chemical Intermediates in Catalysis by TDP

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-

12

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

16

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.

Mechanisms and Structures of Thiamine Biosynthesis

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-

Mechanisms and Structures of Thiamine Biosynthesis

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.

Mechanisms and Structures of Thiamine Biosynthesis

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.

Mechanisms and Structures of Thiamine Biosynthesis

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.

3.

4.

5.

6.

7.

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.

Mechanisms and Structures of Thiamine Biosynthesis 9. 10.

11.

12.

13.

14. 15.

16.

17.

18. 19.

20.

21. 22.

23.

24.

27

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-

28

25. 26. 27. 28.

29.

30.

31.

32.

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

31

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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33

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.

Structure and Function of TPK

35

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.)

Structure and Function of TPK

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

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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.

3.

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.

Structure and Function of TPK 4. 5.

6. 7. 8.

9.

10. 11. 12.

13. 14. 15. 16. 17.

18.

19.

20. 21.

41

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-

42

22.

23.

24. 25.

26. 27.

28.

29. 30. 31. 32.

33.

34. 35.

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

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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 <0.1 lM). Half-maximal incorporation of 32P in rapsyn is observed at 5–28 lM TTP (9), while chloride channels are activated in excised inside-out patches at 1–10 lM (8). Therefore, it might be expected that TTP has no eﬀect under normal physiological conditions. Presently unknown signals would cause a down-regulation of ThTPase activity and/or an up-regulation of TTP synthesis, leading to increased TTP concentrations and phosphorylation of target proteins, leading to a cellular response (Fig. 6). Because TTP is present in all organisms, this may be a very basic mechanism, possibly related to to cell division or protein synthesis.

ACKNOWLEDGMENTS This work was supported by the National Funds for Scientiﬁc Research (FNRS). The authors are Research Associates at the FNRS.

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55

REFERENCES 1. 2. 3. 4.

5. 6. 7. 8.

9.

10.

11. 12.

13. 14.

15. 16.

17.

18.

A Rossi-Fanelli, N Siliprandi, P Fasella. On the presence of triphosphothiamine (TPT) in the liver. Science 116:711–713, 1952. K.-H Kiessling. Thiamine triphosphate in bakers’ yeast. Nature 172:1187–1188, 1953. JR Cooper, JH Pincus. The role of thiamine in nervous tissue. Neurochem Res 4:223–239, 1979. B Minz. Sur la libe´ration de la vitamine B1 par le tronc isole´ du nerf pneumogastrique soumis a´ l’excitation e´lectrique. C R Soc Biol (Paris) 127:1251–1253, 1938. A von Muralt. U¨ber den Nachweis von Aktionssubstanzen der Nervenerregung. Pfu¨lgers Arch 245:604–632, 1942. HP Gurtner. Aneurin und Nervenerregung. Versuche mit 35S markiertem Aneurin and Aneurinantimetaboliten. Helv Physiol Acta, (suppl XI):1–47, 1961. L Bettendorﬀ. Thiamine in excitable tissues: reﬂections on a non-cofactor role. Metab Brain Dis 9:183–209, 1994. L Bettendorﬀ, HA Kolb, E Schoﬀeniels. Thiamine triphosphate activates an anion channel of large unit conductance in neuroblastoma cells. J Membr Biol 136:188–281, 1993. HO Nghieˆm, L Bettendorﬀ, JP Changeux. Speciﬁc phosphorylation of torpedo 43K rapsyn by endogenous kinase(s) with thiamine triphosphate as the phosphate donor. FASEB J 14:543–554, 2000. M Gautam, PG Noakes, J Mudd, M Nichol, GC Chu, JR Sanes, JP Merlie. Failure of postsynaptic specialization to develop at neuromuscular junctions of rapsyn-deﬁcient mice. Nature 377:232–236, 1995. AH West, AN Stock. Histidine kinases and response regulator proteins in twocomponent signaling systems. Trends Biochem Sci 26:369–376, 2001. L Bettendorﬀ, P Wins. Thiamine derivatives in excitable tissues: metabolism, deﬁciency and neurodegenerative diseases. Recent Res Devel Neurochem 2:37– 62, 1999. L Bettendorﬀ. The compartmentation of phosphorylated thiamine derivatives in cultured neuroblastoma cells. Biochim Biophys Acta 1222:7–14, 1994. L Bettendorﬀ, P Wins, M Lesourd. Subcellular localization and compartmentation of thiamine derivatives in rat brain. Biochim Biophys Acta 1222:1–6, 1994. T Eckert, W Mo¨bus. Uber eine ATP: thiaminediphosphat-phosphotransferase— Aktivita¨t im Nervengewebe. H S Z Physiol Chem 338:286–288, 1964. IP Chernikevich, V Luchko, AI Voskoboev, YM Ostrovsky. Puriﬁcation and properties of ATP: thiamine diphosphate phosphotranferase from brewer’s yeast. Biokhimiya 49:899–907, 1984. AI Voskoboev, VS Luchko. Isolation and radiometric determination of rat liver ATP: thiamine diphosphate phosphotransferase activity. Vopr Med Khim 26: 564–568, 1980. K Nishino, Y Itokawa, N Nishino, K Piros, JR Cooper. Enzyme system involved in the synthesis of thiamin triphosphate. I Puriﬁcation and characterization of

56

19.

20.

21.

22.

23.

24.

25.

26.

27.

28. 29. 30.

Bettendorff and Wins protein-bound thiamin diphosphate: ATP phosphoryltransferase. J Biol Chem 258:11871–11878, 1983. Y Egi, S Koyama, H Shikata, K Yamada, T Kawasaki. Content of thiamin phosphate esters in mammalian tissues—an extremely high concentration of thiamin triphosphate in pig skeletal muscle. Biochem Int 12:385–390, 1986. L Bettendorﬀ, C Michel-Cahay, C Grandﬁls, C De Rycker, E Schoﬀeniels. Thiamine triphosphate and membrane-associated thiamine phosphatases in the electric organ of Electrophorus electricus. J Neurochem 49:495–502, 1987. K Miyoshi, Y Egi, T Shioda, T Kawasaki. Evidence for in vivo synthesis of thiamin triphosphate by cytosolic adenylate kinase in chicken skeletal muscle. J Biochem (Tokyo) 108:267–270, 1990. H Shikata, S Koyama, Y Egi, K Yamada, T Kawasaki. Cytosolic adenylate kinase catalyzes the synthesis of thiamin triphosphate from thiamin diphosphate. Biochem Int 18:933–941, 1989. AF Makarchikov, P Wins, F Oerlemans, B Wieringa, T Grisar, L Bettendorﬀ. Adenylate kinase 1 knockout mice have normal thiamine triphosphate levels. Biochim Biophys Acta 1542:117–121, 2002. L Bettendorﬀ, L Weekers, P Wins, E Schoﬀeniels. Injection of sulbutiamine induces an increase in thiamine triphosphate in rat tissues. Biochem Pharmacol 40:2557–2560, 1990. L Bettendorﬀ, M Peeters, P Wins, E Schoﬀeniels. Metabolism of thiamine triphosphate in rat brain: correlation with chloride permeability. J Neurochem 60:423–434, 1993. AF Makarchikov, IP Chernikevich. Puriﬁcation and characterization of thiamine triphosphatase from bovine brain. Biochim Biophys Acta 1117:326– 332, 1992. B Lakaye, AF Makarchikov, AF Antunes, W Zorzi, B Coumans, E De Pauw, P Wins, T Grisar, L Bettendorﬀ. Molecular characterization of a speciﬁc thiamine triphosphatase widely expressed in mammalian tissues. J Biol Chem 277:13771– 13777, 2002. T Nishimune, R Hayashi. Hydrolysis and synthesis of thiamin triphosphate in bacteria. J Nutr Sci Vitaminol (Tokyo) 33:113–127, 1987. J Kyte, RF Doolittle. A simple method for displaying the hydropathic character of a protein. J Mol Biol 157:105–132, 1982. M Hill, A Dupaix, M Nhiri, L Guyen, B Arrio. ATP: ADP phosphotransferase activity, a new characteristic of Catharanthus roseus tonoplasts. FEBS Lett 230:47–50, 1988.

5 How Thiamine Works in Enzymes: Time-Resolved NMR Snapshots of TDP-Dependent Enzymes in Action Kai Tittmann, Ralph Golbik, Kathrin Uhlemann and Gerhard Hu¨bner Martin-Luther-Universita¨t Halle-Wittenberg, Halle/Saale, Germany Ludmila Khailova Russian Academy of Sciences, Moscow, Russia Mulchand S. Patel School of Medicine and Biomedical Sciences, State University of New York at Buﬀalo, Buﬀalo, New York, U.S.A. Frank Jordan Rutgers University, Newark, New Jersey, U.S.A. David M. Chipman Ben-Gurion University of the Negev, Beer-Sheva, Israel Ronald G. Duggleby The University of Queensland, Brisbane, Australia Gunter Schneider Karolinska Institutet, Stockholm, Sweden

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I. INTRODUCTION Thiamindiphosphate (TDP, Fig. 1A), the biologically active form of vitamin B1, is an essential cofactor in biocatalysis in all species, and it mediates important reactions in carbon metabolism, such as the oxidative and nonoxidative decarboxylation of a-keto acids, formation of amino acid precursors, electron transfer reactions, and ketol transfer between sugars (1,2). Studies on the mechanism of activation of enzyme-bound TDP revealed that the 4V-amino group of TDP and a conserved interaction throughout all TDP-dependent enzymes between a glutamate side chain and the N1V nitrogen of the cofactor form a proton relay catalyzing a fast deprotonation at carbon 2 in TDP (3). However, all subsequent steps in catalysis, supposed to proceed via covalent adducts at the C2 position of the thiazolium ring of TDP, could not be observed directly and time-resolved to date. Essentially, even the existence of key intermediates like 2-lactyl-TDP, called ‘‘active pyruvate,’’ could never be proven directly for the enzymic system. Therefore, the intermediate distribution was estimated either by conducting carbon-isotope-eﬀect studies (4,5) or via classical kinetic investigations (6,7). The X-ray structures of many TDP-dependent enzymes were solved and provide structural information on the active sites (8–11); however, the molecular mechanism of single catalytic steps still remains hypothetical. Active site variants of diﬀerent TDP-dependent enzymes showing impaired catalytic constants and altered Michaelis constants could only evidence the general catalytic involvement of side chains, but no compelling conclusions could be deduced with respect to speciﬁc contributions to single steps during substrate turnover. Consequently, only the direct and parallel detection of all covalent intermediates under steady-state or single turnover conditions will provide (1) the validation of the supposed chemical pathway, (2) access to microscopic rate constants of single catalytic steps, (3) thermodynamic quantiﬁcation of

Figure 1 Covalent intermediates in TDP catalysis. (A) Chemical structures and C6V-H 1H NMR ﬁngerprint region of TDP [1] and chemically synthesized intermediates occurring in nonoxidative [3,5] and oxidative enzymic decarboxylation of pyruvate [2,3,5] as well as in enzymic sugar transketolation [4] are shown. The two interfering signals of 2 represent the keto and hydrated forms, respectively (from Ref. 16). Spectra were recorded at pH 0.75 and 25jC. (B) Quantiﬁcation of covalent intermediates in the nonoxidative decarboxylation of pyruvate by ZmPDC and selected enzyme variants during steady-state conditions at substrate saturation. The intermediates were isolated by acid quench and analyzed by 1H NMR at pH 0.75 using the characteristic C6V-H chemical shifts. Their relative populations were determined from the relative integrals of the corresponding signals and used to calculate all microscopic rate constants of the catalytic cycle.

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the stabilization/destabilization of intermediates, and (4) elucidation of mechanistic principles of catalysis at a molecular level (12). Based on the excellent work of Holzer, Krampitz, Kluger, and Frey in the ﬁeld of chemical synthesis and characterization of C2-TDP adducts, we successfully synthesized the supposed key intermediates 2-lactyl-TDP (LTDP), 2-(1-hydroxyethyl)-TDP (HETDP), 2-acetyl-TDP (AcTDP), and 2-(1,2)-dihydroxyethyl-TDP (DHETDP) (Fig. 1A) (13–16). An investigation of the chemical stabilities and spectroscopic properties of those compounds revealed that all chemically synthesized C2 adducts do not undergo decomposition below pH 1 and can be discriminated unequivocally using 1H NMR under these conditions. Particularly, the 1H NMR chemical shifts of the C6VH singlets of the aminopyrimidinium moiety of TDP and the respective TDP adducts are dependent on the chemical nature of the substituent at C2, thus providing a ‘‘ﬁngerprint region’’ for the direct, parallel, and quantitative detection of all C2-derived covalent TDP adducts (y ppm C6V-H: TDP 8.01, LTDP 7.26, HETDP 7.33; AcTDP 7.36, 7.37, 8.6,* DHETDP 7.31) (Fig. 1A). In a series of NMR-based experiments we found that the covalent C2 adducts do not exhibit altered stabilities in intermediate mixtures and in the presence of even excessive amounts of all common native substrates and products of TDP-dependent enzymes. The C6V-H ﬁngerprint region is not obscured by 1H NMR signals from substrates, products, and buﬀers.

II. SETTING UP THE ENZYMIC SYSTEM In order to isolate and quantify covalent TDP adducts in the enzymic system it is possible to apply either the steady-state or the single-turnover approach. Given the diﬃculty that the catalytic constants (kcat) of TDP-dependent enzymes range up to a few hundred per second, the subsequent formation and interconversion of covalent intermediates under single-turnover conditions may not be time resolvable for very fast enzymes using rapid mixing techniques. These limitations can be overcome when the catalytically competent intermediates are coisolated under steady-state conditions. In doing so, the occurrence and distribution of intermediates reﬂect their ‘‘persistent’’ steadystate concentration. The ratio of the concentrations of the intermediates in steady state is determined by the ratio of the microscopic rate constants of their interconversion, which can be straightforwardly transformed into absolute values in consideration of the overall catalytic constant. For example, the

* 2-Acetyl-TDP is in equilibrium with its hydrated form and an internal carbinolamine under these conditions (16).

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catalytic constant of an enzyme involving reversible substrate binding (noncovalent ES complex) and three intramolecular steps K

k2

k3

k4

E þ S ZS ES ! EI1 ! EI2 ! E þ P can be described at substrate saturation as a function of k2, k3, and k4 according to Eq. (1): kcat ¼

k2 k3 k4 k2 k3 þ k2 k4 þ k3 k4

ð1Þ

III. ISOLATION OF COVALENT INTERMEDIATES IN ENZYMIC THIAMINE CATALYSIS Within the framework of our investigations we tried to prove the existence of covalent C2-TDP intermediates in pyruvate decarboxylase (PDC), acetohydroxyacid synthase (AHAS), pyruvate oxidase (POX), pyruvate dehydrogenase (PDH), and transketolase catalysis (TK), respectively. At ﬁrst, enzyme-speciﬁc aspects are considered, followed by a general discussion of basic principles and concepts of covalent thiamine catalysis in enzymes. A. Pyruvate Decarboxylase Pyruvate decarboxylase (PDC, EC 4.1.1.1) is the archetype and is the most intensely studied TDP-dependent enzyme catalyzing the nonoxidative decarboxylation of pyruvate to yield acetaldehyde and carbon dioxide. The minimal catalytic scheme comprises reversible, noncovalent binding of the substrate (Michaelis complex, KS), carbon–carbon bonding (k2) between the C2 of TDP and the carbonyl-carbon of pyruvate to yield LTDP (‘‘active pyruvate’’), subsequent decarboxylation (k3) to the a-carbanion/enamine of HEThDP, and ﬁnally liberation of acetaldehyde (k4) to complete the catalytic cycle: K

k2

k3

E -TDP þ S ZS E -TDP*S ! E -LTDP ! CO2 k4 þE -HETDPðenamine=carbanionÞ þ H2 O ! E -TDP þCH3 CHO þ OH

ð2Þ

In the studies presented here, both the bacterial PDC from Zymomonas mobilis (ZmPDC) and the allosterically regulated enzyme from Saccharomyces cereviseae (ScPDC) were investigated regarding their intermediate distribution in steady state. 1 H NMR spectroscopic analysis of the isolated covalent intermediates of ZmPDC, derived from the acid quench (TCA/HCl) of the working wild-

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type enzyme at steady state after 35 ms reaction time with pyruvate, directly proved the existence of 2-lactyl-TDP and 2-(1-hydroxyethyl)-TDP* as catalytically competent intermediates, since the 1H NMR chemical shifts and spin systems are identical with the chemically synthesized C2-TDP adducts (Fig. 1B). The quantitative intermediate distribution was essentially the same at diﬀerent reaction times, ranging from 20 to 100 ms, thus excluding any presteady-state processes or product inhibition resulting in altered intermediate distributions. Interestingly, the apparent occurrence of TDP in steady state is not due to the dissociation of the cofactor from the enzyme, as evidenced by the retention of full enzymatic activity. Moreover, the quantitative distribution of TDP, LTDP, and HETDP is unchanged at varied pyruvate concentrations of 15–100 mM, clearly pointing to a unimolecular step prior to the decarboxylation of LTDP. Based on the determined steady-state distribution of all intermediates and kcat, the microscopic rate constants of all catalytic steps can be calculated according to Eq. (1) (Table 1). The resulting rate constants show both the decarboxylation (k3) and acetaldehyde release (k4) to be partially rate limiting, as well as a very fast formation of LTDP (k2). A comparable intermediate distribution was determined for the activated ScPDC (Table 1). The steady-state intermediate distribution in both ZmPDC and ScPDC estimated from carbon-isotopeeﬀect studies (4,5) is in excellent agreement with our NMR results. Apparently, the potential of this new method for allowing the parallel and direct observation of all covalent intermediates opens up the way for a detailed mechanistic elucidation of single steps by using rationally designed active-site variants. Therefore, we have determined the steady-state intermediate distribution of various active-site variants of ZmPDC with greatly impaired overall activity. Within the scope of our studies we investigated variants of (1) Glu473 being in a perpendicular orientation to the thiazolium moiety of ThDP, (2) Asp27 and His113, which are in hydrogen-bonding distance to each other in the active-site cleft, and (3) Glu50 interacting with the N1V of the enzyme-bound TDP (Fig. 2). A mutation of Glu473Asp leads to an enzyme variant that is deﬁcient in catalyzing the decarboxylation of LTDP, since the latter is the major populated intermediate in steady state (Fig. 1B). Moreover, the rate of intramolecular substrate binding is also diminished. The rate of C2-H ionization is far from rate limiting; hence carbon–carbon bonding between C2 of TDP and Ca of pyruvate is aﬀected in this variant. Since Glu473 is evidently in-

* The isolated HETDP comprises the fraction of catalytically competent HETDP and the fraction of the a-carbanion/enamine form of HETDP that is instantaneously solvent protonated in the course of acid quench isolation.

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Table 1 Microscopic Rate Constants of Elementary Catalytic Steps (compare Fig. 1B) of ZmPDC Wild Type (wt), ZmPDC Variants, and Activated ScPDC Wild Type

ZmPDC wt ZmPDC wt (D2O) ZmPDC Glu473Asp ZmPDC Asp27Glu ZmPDC His113Lys ZmPDC Glu50Gln ScPDC wt

kcat (s1)

kobs (s1)a H/D exchange

k2 (s1)b C–C bonding

k3 (s1)b CO2 release

k4 (s1)b Acetaldehyde release

F F F F F F F

110 F 19 — 104 F 20 117 F 13 96 F 24 25 F 7 >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

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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.

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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.)

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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).

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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

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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.

5.

6.

7.

8. 9.

10.

11.

12. 13.

A Schellenberger. Sixty years of thiamin diphosphate biochemistry. Biochim Biophys Acta 1385:177–186, 1998. RL Schowen. In: Sinnot M, ed. Comprehensive Biological Catalysis. Vol 2. London: Academic Press, 1998, pp. 217–266. D Kern, G Kern, H Neef, K Tittmann, M Killenberg-Jabs, C Wikner, G Schneider, G Hu¨bner. How thiamin is activated in enzymes. Science 275:67–70, 1997. FJ Alvarez, J Ermer, G Hu¨bner, A Schellenberger, Schowen RL. 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. S Sun, RG Duggleby, RL Schowen. The linkage of catalysis and regulation in enzyme action. Carbon isotope eﬀects, solvent isotope eﬀects and proton inventories for the unregulated pyruvate decarboxylase of Zymomonas mobilis. J Am Chem Soc 117:7317–7322, 1995. CY Huang, AK Chang, PF Nixon, RG Duggleby. Site-directed mutagenesis of the ionizable groups in the active site of Zymomonas mobilis pyruvate decarboxylase: eﬀect on activity and pH dependence. Eur J Biochem 268:3558–3565, 2001. 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. YA Muller, GE Schulz. Structure of the thiamine- and ﬂavin-dependent enzyme pyruvate oxidase. Science 259:965–967, 1993. 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 diphosphatedependent enzyme pyruvate decarboxylase from the yeast Saccharomyces cereviseae at 2.3-A˚ resolution. J Mol Biol 256:590–600, 1996. D Dobritzsch , S Ko¨nig, G Schneider, Lu G. High-resolution crystal structure of pyruvate decarboxylase from Zymomonas mobilis. Implications for substrate activation in pyruvate decarboxylases. J Biol Chem 273:20196–20204, 1998. Y Lindqvist, G Schneider, U Ermler, M Sundstrom. Three-dimensional structure of transketolase, a thiamine diphosphate–dependent enzyme, at 2.5-A˚ resolution. EMBO J 11:2373–2379, 1992. A Fersht. Structure and Mechanism in Protein Science. New York: W. H. Freeman, 1999, pp 216–244. R Kluger, T Smyth. Interaction of pyruvate–thiamin diphosphate adducts with

76

14. 15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

Tittmann et al. pyruvate decarboxylase. Catalysis through closed transition states. J Am Chem Soc 103:1214–1216, 1981. L Krampitz, G Greull, CS Miller, JB Bicking, HR Skeggs, JM Sprague. An active acetaldehyde-thiamin intermediate. J Am Chem Soc 80:5893–5894, 1958. H Holzer, K Beaucamp. Nachweis und Charakterisierung von Zwischenprodukten der Decarboxylierung und Oxydation von Pyruvat: ‘‘aktiviertes Pyruvat’’ und ‘‘aktivierter Acetaldehyd’’. Angew Chem 24:776, 1959. KJ Gruys, CJ Halkides, PA Frey. Synthesis and properties of 2-acetylthiamin pyrophosphate: an enzymatic reaction intermediate. Biochemistry 26:7575– 7585, 1987. M Lobell, DHG Crout. Pyruvate decarboxylase: a molecular modeling study of pyruvate decarboxylation and acyloin formation. J Am Chem Soc 118:1867– 1873, 1996. YG Wu, AK Chang , PF Nixon, W Li, Duggleby RG. Mutagenesis at asp27 of pyruvate decarboxylase from Zymomonas mobilis. Eﬀect on its ability to form acetoin and acetolactate. Eur J Biochem 267:6493–6500, 2000. R Kluger, J Chin, T Smyth. Thiamin-catalyzed decarboxylation of pyruvate. Synthesis and reactivity analysis of the central, elusive intermediate, a-lactylthiamin. J Am Chem Soc 103:884–888, 1981. J Crosby, R Stone, GE Lienhard . Mechanism of thiamine-catalyzed reactions. Decarboxylation of 2-(1-carboxy-1-hydroxyethyl)-3,4-dimethylthiazolium chloride. J Am Chem Soc 92:2891–2900, 1970. 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. 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. K Tittmann, D Proske, M Spinka, S Ghisla, R Rudolph, G Hu¨bner, G Kern. Activation of thiamin diphosphate and FAD in the phosphate-dependent pyruvate oxidase from Lactobacillus plantarum. J Biol Chem 273:12929–12934, 1998. K Tittmann, R Golbik, S Ghisla, G Hu¨bner. Mechanism of elementary catalytic steps of pyruvate oxidase from Lactobacillus plantarum. Biochemistry 39:10747– 10754, 2000. 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 alpha-carbanion of (alpha,beta-dihydroxyethyl)-thiamin diphosphate in the active site of transketolase from Saccharomyces cerevisiae. Proc Natl Acad Sci USA 99:591–595, 2002.

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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REFERENCES 1.

2. 3. 4. 5. 6. 7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

M Pohl. Protein design on pyruvate decarboxylase (PDC) by site-directed mutagenesis. In: T Scheper, ed. Advances in Biochemical Engineering/Biotechnology. Vol. 58. Heidelberg: Springer, Verlag, 1997, pp 15–43. U Scho¨rken, GA Sprenger. Thiamin-dependent enzymes as catalysts in chemoenzymatic syntheses. Biochim Biophys Acta 1385:229–243, 1998. 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 Catal B 6:145–159, 1999. OP Ward, A Singh. Enzymatic asymmetric synthesis by decarboxylases. Curr Opin Biotechnol 11:520–526, 2000. NJ Turner. Applications of transketolases in organic synthesis. Curr Opin Biotechnol 11:527–531, 2000. R Wilcocks, OP Ward, S Collins, NJ Dewdney, Y Hong, E Prosen. Acyloin formation by benzoylformate decarboxylase from Pseudomonas putida. Appl Environm 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, 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. 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. DHG Crout. The chemistry of branched-chain amino acid biosynthesis: stereochemical and mechanistic aspects. In: Z Barak, DM Chipman, V Schloss, eds., Biosynthesis of Branched Chain Amino Acids. Weinheim: Chemie Verlag, 1990, pp 199–242. 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. S Engel, M Vyazmensky, S Geresh, Z Barak, DM Chipman. Acetoxydroxyacid Synthase: A new enzyme for chiral synthesis of R-phenylacetylcarbinol. Biotechnol Bioeng (in press). Z Guo, A Goswami, KD Mirfakhrae, RN Patel. Asymmetric acyloin condensation catalyzed by phenylpyruvate decarboxylase. Tetrahedron: Asymmetry 10:4667–4675, 1999. Z Guo, A Goswami, VB Nanduri, RN Patel. Asymmetric acyloin condensation catalyzed by phenylpyruvate decarboxylase. Part 2: Substrate speciﬁcity and puriﬁcation of the enzyme. Tetrahedron: Asymmetry 12:571–577, 2001. GA Sprenger, U Scho¨rken, T Wiegert, S Grolle, AA de Graaf, SV Taylor, TP

Thiamine-Dependent Enzymes and C–C Bond Formation

17.

18.

19.

20.

21.

22. 23.

24. 25.

26.

27.

28.

29.

89

Begley, S Bringer-Meyer, H Sahm. Identiﬁcation of a thiamine-dependent synthase in Escherichia coli required for the formation of the 1-deoxy-D-xylulose-5-phosphate precursor to isoprenoids, thiamine, and pyridoxol. Proc Natl Acad Sci USA 94:12857–12862, 1997. SV Taylor, LD Vu, TP Begley, U Scho¨rken, S Grolle, GA Sprenger, S BringerMeyer, H Sahm. Chemical and enzymatic synthesis of 1-deoxy-D-xylulose-5phosphate. J Org Chem 63:2375–2377, 1998. F Rohdich, K Kis, A Bacher, W Eisenreich. The non-mevalonate pathway of isoprenoids: genes, enzymes and intermediates. Curr Opin Chem Biol 5:535– 540, 2001. S Hecht, K Kis, W Eisenreich, S Amslinger, J Wungsintaweekul, S Herz, F Rohdich, A Bacher. Enzyme-assisted preparation of isotope-labeled 1-deoxy-Dxylulose 5-phosphate. J Org Chem 66:3948–3952, 2001. 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 1:633–635, 2001. 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. J Koga. Structure and function of indolepyruvate decarboxylase, a key enzyme in indole-3-acetic acid biosynthesis. Biochim Biophys Acta 1249:1–13, 1995. MT Brandl, SE Lindow. Cloning and characterization of a locus encoding an indolepyruvate decarboxylase involved in indole-3-acetic acid synthesis in Erwinia herbicola. Appl Environm Microbiol 62:4121–4128, 1996. F Ama´rita, D Ferna´ndez-Espla´, T Requena, C Pelaez. Conversion of methionine to methional by Lactococcus lactis. FEMS Microbiol Lett 204:189–195, 2001. M Graupner, H Xu, RH White. Identiﬁcation of the gene encoding sulfopyruvate decarboxylase, an enzyme involved in biosynthesis of coenzyme M. J Bacteriol 182:4862–4867, 2000. H Nakashita, K Watanabe, O Hara, T Hidaka, H Seto. Studies on the biosynthesis of bialaphos. Biochemical mechanism of C–P bond formation: discovery of phosphonopyruvate decarboxylase, which catalyzes the formation of phosphonoacetaldehyde from phosphonopyruvate. J Antibiot 50:212–219, 1997. D Schwartz, J Recktenwald, S Pelzer, W Wohlleben. Isolation and characterization of the PEP-phosphomutase and the phosphonopyruvate decarboxylase genes from the phosphinothricin tripeptide producer Streptomyces viridochromogenes Tu¨494. FEMS Microbiol Lett 163:149–157, 1998. R Meganathan, R Bentley. Thiamine pyrophosphate requirement for o-succinylbenzoic acid synthesis in Escherichia coli and evidence for an intermediate. J Bacteriol 153:739–746, 1983. A Weische, M Johanni, E Leistner. Biosynthesis of o-succinylbenzoic acid I: cell-free synthesis of o-succinylbenzoic acid from isochorismic acid in enzyme

90

30.

31.

32.

33.

34. 35. 36.

37.

38.

39.

40.

41. 42.

43.

Mu¨ller and Sprenger preparations from vitamin K–producing bacteria. Arch Biochem Biophys 256: 212–222, 1987. C Palaniappan, V Sharma, MES Hudspeth, R Meganathan. Menaquinone (vitamin K2) biosynthesis: evidence that the Escherichia coli menD gene encodes both 2-succinyl-6-hydroxy-2,4-cyclohexadiene-1-carboxylic acid synthase and a-ketoglutarate decarboxylase activities. J Bacteriol 174:8111–8118, 1992. Palaniappan, H Taber, R Meganathan. Biosynthesis of o-succinylbenzoic acid in Bacillus subtilis: identiﬁcation of menD mutants and evidence against the involvement of the a-ketoglutarate dehydrogenase complex. J Bacteriol 176:2648– 2653, 1994. LS Green, Y Li, DW Emerich, FJ Bergersen, DA Day. Catabolism of aketoglutarate by a sucA mutant of Bradyrhizobium japonicum: evidence for an alternative tricarboxylic acid cycle. J Bacteriol 182:2838–2844, 2000. S Shigeoka, T Onishi, K Maeda, Y Nakano, S Kitaoka. Occurrence of thiamin pyrophosphate–dependent 2-oxoglutarate decarboxylase in mitochondria of Euglena gracilis. FEBS Lett 195:43–47, 1986. Y-Y Chang, JE Cronan, Jr. Conversion of Escherichia coli pyruvate oxidase to an ‘‘a-ketobutyrate oxidase.’’ Biochem J 352:717–724, 2000. G Krakow, SS Barkulis. Conversion of glyoxylate to hydroxypyruvate by extracts of Escherichia coli. Biochim Biophys Acta 21:593–594, 1956. Y-Y Chang, AY Wang, JE Cronan Jr. Molecular cloning, DNA sequencing, and biochemical analyses of Escherichia coli glyoxylate carboligase. J Biol Chem 268:3911–3919, 1993. ZA Janowicz, MR Eckart, C Drewke, RO Roggenkamp, CP Hollenberg. Cloning and characterization of the DAS gene encoding the major methanol assimilatory enzyme from the methylotrophic yeast Hansenula polymorpha. Nucl Acids Res 13:3043–3062, 1985. YT Ro, CY Eom, T Song, JW Cho, YM Kim. Dihydroxyacetone synthase from a methanol-utilizing carboxydobacterium, Acinetobacter sp. strain JC1 DSM 3803. J Bacteriol 179:6041–6047, 1997. M Miyazaki, M Shibue, K Ogino, H Nakamura, H Maeda. Enzymatic synthesis of pyruvic acid from acetaldehyde and carbon dioxide. Chem Commun 18:1800– 1801, 2001. V Foulon, VD Antonenkov, K Croes, E Waelkens, GP Mannaerts, PP van Veldhoven, M Casteels. Puriﬁcation, molecular cloning, and expression of 2hydroxyphytanoyl-CoA lyase, a peroxisomal thiamine pyrophosphate–dependent enzyme that catalyzes the carbon–carbon bond cleavage during aoxidation of 3-methyl-branched fatty acids. Proc Natl Acad Sci USA 96:10039– 10044, 1999. AJ Baetz, MJ Allison. Puriﬁcation and characterization of oxalyl-coenzyme A decarboxylase from Oxalobacter formigenes. J Bacteriol 171:2605–2608, 1989. H-Y Lung, AL Baetz, AB Peck. Molecular cloning, DNA sequence, and gene expression of the oxalyl-coenzyme A decarboxylase gene, oxc, from the bacterium Oxalobacter formigenes. J Bacteriol 176:2468–2472, 1994. S Ui, T Hosaka, K Mizutani, A Mimura. Puriﬁcation and properties of acetyl-

Thiamine-Dependent Enzymes and C–C Bond Formation

44.

45.

46.

47.

48.

49.

50.

51.

52. 53.

91

acetoin synthase from Bacillus sp. YUF-4. Biosci Biotechnol Biochem 62:795– 797, 1998. S Ui, T Hosaka, K Mizutani, T Ohtsuki, A Mimura. Acetylacetoin synthase as a marker enzyme for detecting the 2,3-butanediol cycle. J Biosci Bioeng 93:248– 251, 2002. H Chen, Z Guo, H-W Liu. Biosynthesis of Yersiniose: attachment of the twocarbon branched-chain is catalyzed by a thiamine pyrophosphate–dependent ﬂavoprotein. J Am Chem Soc 120:11796–11797, 1998. Z Zhao, H-W Liu. Synthesis of a deoxysugar dinucleotide containing an exodiﬂuoromethylene moiety as a mechanistic probe for studying enzymes involved in unusual sugar biosynthesis. J Org Chem 66:6810–6815, 2001. J Guo, JW Frost. Biosynthesis of 1-deoxy-1-imino-D-erythrose-4-phosphate: a deﬁning metabolite in the aminoshikimate pathway. J Am Chem Soc 124:528– 529, 2002. J Castells, F Lo´pez-Calahorra, M Bassedas, P Urrios. A new thiazolium salt– catalyzed synthesis of a-aminoketones from aldehydes and iminium salts. Synthesis 314–315, 1988. AR Katrizky, D Cheng, RP Musgrave. a-Amino ketone synthesis from aldehydes and N-(a-dialkylaminoalkyl)benzotriazoles. Heterocycles 42:273–280, 1996. JA Murry, DE Frantz, A Soheili, R Tilyer, EJJ Grabowski, PJ Reider. Synthesis of a-amido ketones via organic catalysis: thiazolium-catalyzed crosscoupling of aldehydes with acylimines. J Am Chem Soc 123:9696–9697, 2001. N Khaleeli, R Li, CA Townsend. Origin of the h-lactam carbons in clavulanic acid from an unusual thiamine pyrophosphate–mediated reaction. J Am Chem Soc 121:9223–9224, 1999. M Goldberg, JM Fessenden, E Racker. Phosphoketolase. Meth Enzymol 9:515– 520, 1996. T Kitazume, N Ishikawa. Introduction of center of chirality into ﬂuorocompounds by microbial transformation of 2,2,2-triﬂuoroethanol. Chem Lett 1815– 1818, 1984.

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

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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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4. 5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

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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.

112

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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

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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.

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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.

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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 Vicun˜a (22). They showed that this strain can grow on benzoin as a sole carbon and energy source, due to the ability of BAL

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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

128

5.

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 Vicun˜a. 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 Vicun˜a. 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.

REFERENCES 1. 2.

3.

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

BFD: Lessons in Enzymology

4. 5. 6.

7.

8.

9. 10.

11.

12.

13.

14.

15.

141

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.

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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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REFERENCES 1.

2.

3. 4.

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

GA Kochetov. Transketolase from yeast, rat liver and pig liver. In: Willis A. Wood, ed. Methods in Enzymology. New York: Academic Press, 1982, v. 90, pp. 209–223. 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. 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. M Sundstro¨m, Y Lindqvist, G Schneider. Three-dimensional structure of apotransketolase. Flexible loops at the active site enable cofactor binding. FEBS Lett 313:229–231, 1992. G Schneider, Y Lindqvist. Crystallography and mutagenesis of transketolase: mechanistic implications for enzymatic thiamin catalysis. Biochim Biophys Acta 1385:387–398, 1998. U Nilsson, Y Lindqvist, R Klu¨ger, G Schneider. Crystal structure of transketolase in complex with thiamin thiazolone diphosphate, an analogue of the reaction intermediate, at 2.3-A˚ resolution. FEBS Lett 326:145–148, 1993. 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. U Nilsson, L Meshalkina, Y Lindqvist, G Schneider. Examination of substrate binding in thiamine diphosphate–dependent transketolase by protein crystallography and site-directed mutagenesis. J Biol Chem 272:1864–1869, 1997. 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. PNAS 99:591–595, 2002. C Wikner, L Meshalkina, U Nilsson, M Nikkola, Y Lindqvist, M Sundstro¨m, G Schneider. Analysis of an invariant cofactor–protein interaction in thiamin diphosphate–dependent enzymes by site-directed mutagenesis. J Biol Chem 269:32144–32150, 1994. C Wikner, L Meshalkina, U Nilsson, S Backstro¨m, Y Lindqvist, G Schneider. His103 in yeast transketolase is required for substrate recognition and catalysis. Eur J Biochem 233:750–755, 1995. L Meshalkina, U Nilsson, C Wikner, T Kostikova, G Schneider. Examination of the thiamin diphosphate binding site in yeast transketolase by site-directed mutagenesis. Eur J Biochem 244:646–652, 1997. C Wikner, U Nilsson, L Meshalkina, C Udekwu, Y Lindqvist, G Schneider. Identiﬁcation of catalytically important residues in yeast transketolase. Biochemistry 36:15643–15649, 1997. U Nilsson, L Hecquet, T Geﬄaut, C Guerard, G Schneider. Asp477 is a determinant of the enantioselectivity in yeast transketolase. FEBS Lett 42:49– 52, 1998.

156

Kovina et al.

15. M Sundstro¨m, Y Lindqvist, G Schneider, U Hellman, H Ronne. Yeast TKL1 gene encodes a transketolase that is required for eﬃcient glycolysis and biosynthesis of aromatic amino acids. J Biol Chem 268:24346–24352, 1993. 16. SW Cavalieri, KE Neet, HZ Sable. Enzymes of pentose biosynthesis. The quaternary structure and reacting form of transketolase from baker’s yeast. Arch Biochem Biophys 171:527–532, 1975. 17. GA Kochetov, LE Meshalkina, RA Usmanov. The number of active sites in a molecule of transketolase. Biochem Biophys Res Commun 69:839–843, 1976. 18. GA Kochetov, PP Philippov. Calcium: cofactor of transketolase from backer’s yeast. Biochem Biophys Res Commun 38:930–933, 1970. 19. PC Heinrich, H Steﬀen, P Janser, O Wiss. Studies on the reconstitution of apotransketolase with thiamine diphosphate and analogs of the coenzyme. Eur J Biochem 30:533–541, 1972. 20. Y Mu¨ller, Y Lindqvist, W Furey, G Schulz, F Jordan, G Schneider. A thiamine diphosphate binding fold revealed by comparison of the crystal structures of transketolase, pyruvate oxidase and pyruvate decarboxylase. Structure 1:95– 103, 1993. 21. F Dyda, W Furey, S Swaminathan, M Sax, B Farrenkopf, F Jordan. Catalytic centers in the thiamin diphosphate enzyme pyruvate decarboxylase at 2.4-A˚ resolution. Biochemistry 32:6165–6170, 1993. 22. YA Mu¨ller, GE Schulz. Structure of the thiamin- and ﬂavin-dependent enzyme pyruvate oxidase. Science 259:965–967, 1993. 23. M Killenberg-Jabs, S Ko¨nig, I Eberhard, 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. 24. GA Kochetov, RA Usmanov, VP Merzlov. Thiamin pyrophosphate–induced change of the optical activity of baker’s yeast transketolase. FEBS Lett 9:265– 266, 1970. 25. CP Heinrich, K Noack, O Wiss. A circular dichroism study of transketolase from baker’s yeast. Biochem Biophys Res Commun 44:275–279, 1971. 26. LE Meshalkina, GA Kochetov. The functional identity of the active centers of transketolase. Biochim Biophys Acta 571:218–223, 1979. 27. MV Kovina, VA Selivanov, NV Kochevova, GA Kochetov. Kinetic mechanism of active-site non-equivalence in transketolase. FEBS Lett 418:11–14, 1997. 28. RA Usmanov, GA Kochetov. Interaction of baker’s yeast transketolase with substrates. Biochem Internat 5:727–734, 1982. 29. IA Bykova, ON Solovjeva, LE Meshalkina, MV Kovina, GA Kochetov. Onesubstrate transketolase-catalyzed reaction. Biochem Biophys Res Commun 280:845–847, 2001. 30. ON Solovjeva, IA Bykova, LE Meshalkina, MV Kovina, GA Kochetov. Cleaving of ketosubstrates by transketolase and the nature of the products formed. Biochemistry (Moscow) 66:1144–1149, 2001. 31. E Fiedler, R Golbik, G Schneider, K Tittmann, H Neef, S Ko¨nig, G Hu¨bner. Examination of donor substrate conversion in yeast transketolase. J Biol Chem 276:16051–16058, 2001.

Induced Absorption Band of Holotransketolase

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32. GA Kochetov, RA Usmanov, AT Mevkh. The role of the charge transfer complex in the transketolase catalyzed reaction. Biochem Biophys Res Commun 54: 1619–1626, 1973. 33. LE Meshalkina, C Wikner, U Nilsson, Y Lindqvist, G Schneider. CD spectra of recombinant wild-type and mutant transketolase. In: H Bisswanger, A Schellenberger, eds. Biochemistry and Physiology of Thiamin Diphosphate Enzymes. A.u.C. Intemann, Wissenschaftlicher Verlag, Prien, On Chiemsee, Germany, 1996, pp. 532–542. 34. G Schneider, Y Lindqvist. Enzymatic thiamine catalysis: mechanistic implication from the three-dimensional structure of transketolase. Bioorg Chem 21:109–117, 1993. 35. D Kern, G Kern, H Neef, K Tittmann, M Killenberg-Jabs, C Wikner, G Hu¨bner, G Hu¨bner. How thiamine diphosphate is activated in enzymes. Science 275:67–70, 1997. 36. W Furey, P Arjunan, L Chen, M Sax, F Guo, F Jordan. Structure–functional relationship and ﬂexible tetramer assembly in pyruvate decarboxylase revealed by analysis of crystal structures. Biochim Biophys Acta 1385:253–270, 1998. 37. YM Ostrovsky, II Stepuro, A Schellenberger, G Hu¨bner. The electronic spectrum of thiamin and properties of molecule of vitamin in the hydrophobic surrounding. Biokhimiya (in Russian) 36:1222–1227, 1971. 38. A Schellenberger. Sixty years of thiamine diphosphate biochemistry. Biochim Biophys Acta 1385:177–186, 1998. 39. DE Metzler. Thiamine coenzymes. In: PD Boyer, H Lardy, K Myrba¨ck, eds. The Enzymes. New York: Academic Press, 1960, v. 2, pp. 295–337. 40. R Friedemann, H Neef. Theoretical studies on the electronic and energetic properties of the aminopyrimidine part of thiamin diphosphate. Biochim Biophys Acta 1385:245–250, 1998. 41. MG Pustynnikov, H Neef, RA Usmanov, A Schellenberger, GA Kochetov. Functional groups of thiamine pyrophosphate in holotransketolase. Biokhimiya (in Russian) 51:1003–1016, 1986. 42. R Golbik, H Neef, G Hu¨bner, S Ko¨nig, B Seliger, L Meshalkina, GA Kochetov, A Schellenberger. Function of the aminopyrimidine part in thiamine pyrophosphate enzymes. Bioorg Chem 19:10–17, 1991. 43. TL Gilchrist. Heterocyclic Chemistry. 2nd ed. New York: Longman Scientiﬁc and Technical, Co-published in the U.S. with Wiley, 1982. 44. K Inuzuka, A Fujimoto. The amino-imino tautomerization of the 2-aminopyridine-acetic acid system in isooctane. Bull Chem Soc Jpn 63:971–975, 1990. 45. W Pietrzycki, J Sepiol, P Tomasik, L Brzozka. Tautomerism and rotamerism in 2-methylamino-, 2-anilino-, 2-acetamido-, and benzamido-pyridines. Bull Soc Chim Belg 102:709–717, 1993.

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 <3.1 A˚) to neighboring residues are indicated by dashed lines.

oxygen interacts through hydrogen bonds with the side chain of His103 and a water molecule, which in turn is anchored to the protein by hydrogen bonds to the side chain of His69 and main chain atoms of residues His69 and Gly116. The interaction with the invariant residue His103 is important for two reasons. It is a key feature for the recognition of the h-hydroxyl group, i.e., the discrimination between hydroxypyruvate and pyruvate. Indeed, mutagenesis data for this amino acid indicated a major role in binding of the donor substrate but not of the acceptor substrate (24). Furthermore, the His103Ala mutant is severely impaired in the formation of the a-carbanion intermediate, consistent with the stabilizing interaction seen in the crystal structure of this intermediate. The a-hydroxyl oxygen forms a hydrogen bond to the side chain of His481. This hydrogen bond also contributes to interactions with the donor substrate, since its replacement with glutamine, serine, or alanine led to signiﬁcantly increased Km values for the donor substrate but only moderate

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decrease in kcat and no changes for the Km values for the acceptor substrate (25). This situation appears to be diﬀerent in mammalian transketolase, where this residue is replaced by glutamine. Site-directed mutagenesis of this glutamine residue in human transketolase only led to minor changes in the kinetic parameters (26). E. Near-UV CD Spectra Reflect Formation of the DHETDP Intermediate The reaction was initiated by soaking of the small donor substrate into crystals of the holo-enzyme. In spite of the time required for diﬀusion of the substrate into the active site of the enzyme, the intermediate is already observed with high occupancy in the 30-s data set, indicating that the catalytic steps leading to the a-carbanion must also occur at signiﬁcant rates in the crystal. The lack of any electron density for the carboxyl group of hydroxypyruvate suggests further that the decarboxylation step is fast as compared to the decomposition of the a-carbanion into TDP and glycolaldehyde. This in turn supports the proposal (19) that the spectroscopic changes in the absorption and CD spectra around 300–320 nm are due to the presence of the a-carbanion/enamine of DHETDP at the active site of the enzyme. F. Conformational Changes Are Not Required for Intermediate Formation The steps from the binding of donor substrate to the formation of the acarbanion intermediate can occur within the crystal lattice, and we do not observe any large conformational changes in the enzyme during these catalytic steps, which represent the ﬁrst half of the transketolase reaction. Structural changes upon formation of the enzyme intermediate, which had been postulated based on molecular dynamics simulations (27), are thus not supported by the present crystal structure analysis. The second part of the reaction, transfer of the glycolaldehyde moiety onto an acceptor substrate, is in principal the reversal of the ﬁrst catalytic steps. Together with the observation that binding of the acceptor substrate erythrose-4-phosphate to holotransketolase does not induce any large-scale conformational changes in the enzyme (28), it appears unlikely that catalysis by transketolase is associated with any signiﬁcant structural changes. Conformational transitions that might be invoked to occur transiently, i.e., escape detection by crystallography, must also be limited in size, since the crystal lattice is not disturbed upon addition of the donor substrate HPA. The reaction intermediate accumulates to similar high occupancies in the two active sites of transketolase, as indicated by the electron density maps and the B-factors for the a,h dihydroxyethyl moiety of the bound

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DHETDP. The active sites in the dimer are thus independent of each other, and our results are not compatible with proposals of half-of-the-site reactivity of transketolase (29). G. Implications for Other TDP-Dependent Enzymes In transketolase, the carbanion/enamine acts as nucleophile in the second step of the reaction. The enzyme therefore has to avoid protonation of the carbanion/enamine. This is diﬀerent from the situation in, for instance, pyruvate decarboxylase, where protonation of the carbanion/enamine is part of the catalytic mechanism. These mechanistic diﬀerences are reﬂected in the activesite topology of the two enzymes. In pyruvate decarboxylase a residue (Glu477) is suitably positioned to participate in proton transfer (21,30), while in transketolase no such amino acid is found close to the Ca atom of the intermediate. The conformation of the carbanion/enamine intermediate with respect to the thiazolium ring, the Ca, Ch, and the Ca oxygen atoms of the a,hdihydroxyethyl moiety observed in transketolase is very likely common to other TDP-dependent enzymes. For instance, it agrees well with the conformation of the hydroxyethyl TDP in pyruvate decarboxylase derived from modeling studies (21). Spectroscopic data had indicated a signiﬁcant enamine character of the corresponding intermediates in pyruvate decarboxylase (31) and benzoylformate decarboxylase (32), and it thus appears that this is a

Figure 5 Comparison of the structures of the thiazolium ring in the reaction intermediates in transketolase (left) (13) and pyruvate:ferredoxin oxidoreductase (right) (33).

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common characteristic feature for these intermediates in TDP-dependent enzymes. While the last statement may hold for many TDP-dependent enzymes, it is not true for all. The recent crystal structure determination of the complex of pyruvate:ferredoxin oxidoreductase with the free-radical intermediate hydroxyethylidene TDP (33) shows signiﬁcant diﬀerences in the structure of the thiazolium ring, when compared to the a-carbanion/enamine intermediate in transketolase. While in the latter, the thiazolium ring is planar, it is clearly bent in pyruvate:ferredoxin oxidoreductase, indicating signiﬁcant loss of aromaticity (Fig. 5). The structural diﬀerences between the key reaction intermediates of these two TDP-dependent enzymes reﬂect the signiﬁcant variations in their chemistry: Transketolase catalyzes nonoxidative ketol transfer, whereas pyruvate:ferredoxin oxidoreductase uses oxidative chemistry to produce acetyl-CoA from pyruvate and coenzyme A. The crystal structures of the two key intermediates in transketolase and pyruvate:ferredoxin oxidoreductase thus highlight the catalytic versatility of TDP.

ACKNOWLEDGMENTS We gratefully acknowledge access to synchrotron radiation at beam line BW7b, EMBL outstation, DESY, Hamburg. E. F. was supported by grants of the Graduiertenfo¨derung of Sachsen-Anhalt. The work was supported by the Deutsche Forschungsgemeinschaft, the Fonds der Chemischen Industrie, and the Swedish Research Council.

REFERENCES 1.

2. 3.

4.

5. 6. 7.

Sundstro¨m M, Lindqvist Y, Schneider G, Hellman U, Ronne H. Yeast TKL1 gene encodes a transketolase that is required for eﬃcient glycolysis and biosynthesis of aromatic amino acids. J Biol Chem 268:24346–24352, 1993. Sprenger GA, Scho¨rken U, Sprenger G, Sahm H. Transketolase A of Escherichia coli K12. Eur J Biochem 230:525–532, 1995. McCool BA, Plonk SG, Martin PR, Singleton C. Cloning of human transketolase cDNAs and comparison of the nucleotide sequence of the coding region in Wernicke–Korsakoﬀ and Non-Wernicke–Korsakoﬀ individuals. J Biol Chem 268:1397–1404, 1993. Lindqvist Y, Schneider G, Ermler U, Sundstro¨m M. Three-dimensional structure of transketolase, a thiamine pyrophosphate–containing enzyme at 2.5-A˚ resolution. EMBO J 11:2373–2379, 1992. Nikkola M, Lindqvist Y, Schneider G. J Mol Biol 238:387–404, 1994. Schneider G, Lindqvist Y. Bioorg Chem 21:109–117, 1993. Muller Y, Lindqvist Y, Furey W, Schulz GE, Jordan F, Schneider G. The thi-

170

8. 9.

10. 11.

12.

13.

14.

15.

16. 17.

18.

19.

20. 21.

Sandalova et al. amin diphosphate binding fold: comparison of the crystal structures of transketolase, pyruvate oxidase and pyruvate decarboxylase. Structure 1:95–103, 1993. Schellenberger A. Sixty years of thiamin diphosphate biochemistry. Biochem Biophys Acta 1385:177–186, 1998. Jordan F. 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. Kluger R. Thiamin diphosphate: a mechanistic update on enzymic and nonenzymic catalysis of decarboxylation. Chem Rev 87:863–876, 1987. Gubler CJ. Physiological functions and mechanism of action of transketolase. In: H Bisswanger, J Ulrich, eds. Biochemistry and physiology of thiamin diphosphate dependent enzymes. Weinheim, Germany: VCH Verlagsgesellschaft, 1991, pp. 311–321. Fiedler E, Golbik R, Schneider G, Tittmann K, Neef H, Ko¨nig S, Ho¨bner G. Examination of donor–substrate conversion in yeast transketolase. J Biol Chem 276:16051–16058, 2001. Fiedler E, Thorell S, Sandalova T, Golbik R, Konig S, Schneider G. Snapshot of a key intermediate in enzymatic thiamin catalysis: crystal structure of the a-carbanion of (a,h-dihydroxyethyl)-thiamin diphosphate in the active site of transketolase from Saccharomyces cerevisiae. Proc Natl Acad Sci USA 99:591– 595, 2002. Wikner C, Meshalkina L, Nilsson U, Nikkola M, Lindqvist Y, Sundstro¨m M, Schneider G. Analysis of an invariant cofactor–protein interaction in thiamindependent enzymes by site-directed mutagenesis. J Biol Chem 269:32144–32150, 1994. Kochetov GA, Usmanov RA, Mevkh AT. The role of the charge transfer complex in the transketolase-catalyzed reaction. Biochem Biophys Res Commun 54:1619–1626, 1973. Schneider G, Sundstro¨m M, Lindqvist Y. Preliminary Crystallographic data for transketolase from yeast. J Biol Chem 264:21619–21620, 1989. Jones TA, Zou J-Y, Cowan SW, Kjeldgaard M. Improved methods for building protein models in electron density maps and location of errors in these models. Acta Crystallogr D 47:110–119, 1991. Bru¨nger AT, Adams PD, Clore GM, DeLano WL, Gros P, Grosse-Kunstleve RW, Jiang J-S, Kuszewski J, Nilges M, Pannu NS, Read RJ, Rice LM, Simonson T, Warren GL. Crystallography & NMR System: A new software suite for macromolecular structure determination. Acta Crystallogr D 54:905–921, 1998. Kochetov GA, Usmanov RA, Merzlov VP. Thiaminpyrophosphate-induced changes in the optical activity of baker’s yeast transketolase. FEBS Lett 9:265– 266, 1970. Heinrich CP, Noack K, Wiss O. A circular dichroism study of transketolase from baker’s yeast. Biochem Biophys Res Commun 44:275–279, 1971. Lobell M, Crout DHG. Pyruvate decarboxylase: a molecular modelling study of

a-Carbanion/Enamine Intermediate in Transketolase

22.

23.

24.

25.

26.

27.

28.

29. 30.

31.

32.

33.

171

pruvate decarboxylation and acyloin formation. J Am Chem Soc 118:1867–1873, 1996. Golbik R, Neef H, Hu¨bner G, Ko¨nig S, Seliger B, Meshalkina M, Kochetov GA, Schellenberger A. Function of the aminopyrimidine part in thiamine pyrophosphate enzymes. Bioorg Chem 19:10–17, 1991. Kern D, Kern G, Neef H, Tittmann K, Killenberg-Jabs M, Wikner C, Schneider G, Hu¨bner G. How thiamin diphosphate is activated in enzymes. Science 275:67– 70, 1997. Wikner C, Meshalkina L, Nilsson U, Ba¨ckstro¨m S, Lindqvist Y, Schneider G. His103 in yeast transketolase is required for substrate recognition and catalysis. Eur J Biochem 233:750–755, 1995. Wikner C, Nilsson U, Meshalkina L, Udekwu C, Lindqvist Y, Schneider G. Identiﬁcation of catalytically important residues in yeast transketolase. Biochemistry 36:11643–11649, 1997. Singleton CK, Wang JL, Shan L, Martin PR. Conserved residues are functionally distinct within transketolases of diﬀerent species. Biochemistry 35: 15865–15869, 1996. Kovina MV, Tikhonova OV, Soloveva ON, Bykova IA, Ivanov AS, Kochetov GA. Inﬂuence of transketolase substrates on its conformation. Biochem Biophys Res Commun 27:968–972, 2000. Nilsson U, Meshalkina L, Lindqvist Y, Schneider G. Examination of substrate binding in thiamin diphosphate–dependent transketolase by protein crystallography and site-directed mutagenesis. J Biol Chem 272:1864–1869, 1997. Kovina MV, Kochetov GA. Cooperativity and ﬂexibility of active sites in homodimeric transketolase. FEBS Lett 440:81–84, 1998. Dyda F, Furey W, Swaminathan S, Sax M, Farrenopf B, Jordan F. Catalytic centers in the thiamin diphosphate–dependent enzyme pyruvate decarboxylase at 2.4-A˚ resolution. Biochemistry 32:6165–6170, 1993. Zeng X, Chung MH, Jordan F. Direct observation of the kinetic fate of a thiamin diphosphate–bound enamine intermediate on brewer’s yeast pyruvate decarboxylase. Kinetic and regiospeciﬁc consequences of allosteric activation. J Am Chem Soc 113:5842–5849, 1991. Barletta GL, Yu Z, Huskey WP, Jordan F. Kinetics of C(2a)-Proton abstraction from 2-benzylthiazolium salts leading to enamines relevant to catalysis by thiamin-dependent enzymes. J Am Chem Soc 119:2356–2362, 1997. Chabriere E, Vernede X, Guigliarelli B, Charon MH, Hatchikian EC, FontecillaCamps JC. Crystal structure of the free-radical intermediate of pyruvate:ferredoxin oxidoreductase. Science 294:2559–2563, 2001.

12 Yeast Pyruvate Decarboxylase: New Features of the Structure and Mechanism Frank Jordan, Min Liu, Eduard Sergienko, and Zhen Zhang Rutgers University, Newark, New Jersey, U.S.A. Andrew Brunskill, Palaniappa Arjunan, and William Furey Veterans Aﬀairs Medical Center and the University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, U.S.A.

I. INTRODUCTION The structure of yeast pyruvate decarboxylase (YPDC, EC, 4.1.1.1; see Scheme 1 for mechanism), an enzyme that has long served as the ‘‘simplest’’ prototype of thiamine diphosphate (TDP, the vitamin B1 coenzyme)–dependent enzymes, was solved from the strain Saccharomyces uvarum (1) and later from the strain Saccharomyces cerevisiae (2). More recently, the latter structure was redetermined in the presence of the substrate activator surrogate pyruvamide (3). The goal of this chapter is to point out the crucial novel ﬁndings from the authors’ laboratories; general reviews of the mechanistic aspects of TDP enzymes and related models have been published by one of the authors (4,5). 173

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Scheme 1

II. OVERVIEW OF THE YPDC STRUCTURE A. The Monomer The crystals of S. uvarum a4 YPDC ﬁrst studied were monoclinic in space group C2 and contained two complete YPDC monomers in the asymmetric unit. About 95% of the total structure was ﬁtted and reﬁned using X-ray data to 2.4 A˚. The model also included TDP and Mg2+ cofactors as well as water molecules associated with the cofactors. A detailed report on the S. uvarum structure and structure determination was published (1), as was one on the S. cerevisiae structure (2). In both structures each YPDC subunit consists of a single polypeptide chain that folds into three distinct and approximately equal-sized domains named a, h, and g, according to their consecutive locations along the polypeptide chain. All three domains have an a/h topology (Fig. 1). The a and g domains have the same basic fold, with six parallel h strands ﬂanked on both sides by a helices. No interpretable electron density was found for residues 106–113 in the a domain, which are disordered and constitute an exposed loop. The central h domain contains a sevenstranded mixed sheet, with ﬁve strands parallel and the others antiparallel. In the h domain there is no interpretable electron density for residues 292–301, which serve as a crossover connection between two parallel strands. The three domains are centered roughly around the vertices of an equilateral triangle with an edge of about 32 A˚ (Fig. 1). An evident cavity is formed at the center of the triangle and thus between all three domains. The h domain was found

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Figure 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). (See color insert.)

to be much more ﬂexible than either the a or g domains, as indicated by both its larger mean B value (20.5 vs. 12.5 and 14.1 A˚2, respectively) and its larger mean deviation when comparing the corresponding domain pairs in the two monomers present in the asymmetric unit (1.0 vs. 0.29 and 0.38 A˚, respectively). Residues involved in TDP binding are among those with the lowest B values, consistent with a tight binding cofactor. B. The Dimer The two monomers within the crystallographic asymmetric unit are tightly packed to form a dimer with approximate twofold symmetry. Within the dimer, monomer–monomer contacts are formed exclusively between the a and g domains. The TDP binding sites lie at the monomer–monomer interface, with the a domain of monomer 1 and the g domain of monomer 2 binding one TDP and, conversely, the a domain of monomer 2 and the g domain of monomer 1 binding a twofold related TDP. A view of the YPDC dimer is shown in Figure 2. This mode of cofactor binding makes it apparent

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Figure 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). (See color insert.)

that the dimer is the minimal functional unit, since two monomers are required to create the active-site environment, with the approximate twofold symmetry generating two such sites. The disordered regions in the a and h domains from the diﬀerent monomers would be close to each other in space if ordered, due to monomer–monomer interactions in the dimer. C. Varying Tetrameric Assemblies of YPDC To date several distinct tetrameric assemblies of YPDC have been observed, and in each the overall structure of the tightly associated, cofactor-binding dimers remains largely unchanged. It is believed that changes in tetrameric assembly are associated with the substrate activation process. Prior to a discussion of factors that tend to aﬀect the tetrameric assembly of YPDC, a brief description of each tetramer type will be given. In the original C2 monoclinic form of YPDC, a rotation about the crystallographic twofold axis generates another dimer to form a complete YPDC tetramer. Dimer–dimer contacts within the tetramer are conﬁned almost exclusively to the h domains, with the last antiparallel strand in each of these domains interacting reciprocally with the corresponding strand in another h domain. A view of the tetramer is shown in Figure 3. This form generates two large continuous 14 strand h sheets, however, the domaindomain interface between the last strands utilizes only two hydrogen bonds

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Figure 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 space-ﬁlling 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). (See color insert.)

and the ﬁnal strands are also close to perpendicular as shown in the inset to Figure 3. The resulting tetramer is rather loosely associated and leaves the center largely open and accessible to solvent. Given the tight association of the dimers to form the active sites and the loose association between dimers to form the tetramer, the entire tetramer is best described as a dimer of dimers, with approximate 222 symmetry. This assembly type shall be referred to as Form A. To study the structural implications of substrate activation, YPDC was crystallized in the presence of eﬀector molecules, including the substrate surrogate activators ketomalonate (6,7) and pyruvamide (3,8), inhibitors such as HgCl2 and p—ClC6H4—CHjCHCOCOOH, and a number of point mu-

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tants. With ketomalonate a new mode of tetrameric assembly was found in certain crystal forms. A similar tetrameric assembly was later seen when crystallized with pyruvamide, despite the fact that the crystal space group was diﬀerent (C2 for the pyruvamide with a dimer in the asymmetric unit, and P21 for the ketomalonate structure with a tetramer in the asymmetric unit). The major change in this tetrameric assembly was originally described as a rotation of one dimer by roughly 24j about its own internal twofold axis, followed by a translation by about 8 A˚ toward the tetramer center of mass. This assembly type will be called Form B. The end result is a much more compact tetramer with a completely diﬀerent set of dimer–dimer interactions than that observed in Form A. A view of Form B is shown in Figure 4, and a

Figure 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 balland-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). (See color insert.)

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least squares superposition of the ketomalonate- and pyruvamide-containing structures is shown in Figure 5. In Form B a reciprocal hydrogen bond pair remains between h domains on one side of the tetramer but is absent on the other. Thus, in essence the change from A to B involves a pivoting of one dimer relative to the other, with the pivot point being a pair of h-strand hydrogen bonds between T320 main-chain atoms in two subunits. In addition to retaining these hydrogen bonds, a new contact surface is formed with interactions involving both the a and h domains, including some residues that were disordered in Form A. The overall structure of the monomers remains largely unchanged, although there is a rotation of about 8j of the h domain relative to the Form A structure. More notable is the ordering in two of the subunits of certain regions that are disordered in Form A (residues 106–113 and 292–303). Apart from being involved in the new dimer–dimer interface, these new ordered regions are close two of the TDP active sites.

Figure 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). (See color insert.)

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D. Active-Site Environment In both A and B forms of YPDC, the TDP cofactors are situated at the subunit–subunit interface of each tightly associated dimer. The cofactorbinding site from Form A is shown in Figure 6. The diphosphate end of the cofactor is bound exclusively to the g domain of one subunit and the 4Vaminopyrimidine end is bound to this g domain and also to the a domain of the other subunit. Binding of the diphosphate end is achieved by bonding, via an oxygen atom from each of the a and h phosphoryl groups, to an octahedrally coordinated Mg2+, which is also bonded to the side-chain oxygen atoms of D444 and N471, to the main-chain oxygen of G473, and to a water molecule that is bonded to the main-chain oxygen of L469. The residues D444 and N471 are part of a conserved 30-residue stretch previously identiﬁed to be common to all TDP-dependent enzymes (9), and demonstration of its role in YPDC (1,2), transketolase (TK, ref. 10), and pyruvate oxidase (POX, ref. 11), conﬁrms its importance in anchoring the TDP cofactor to the protein. The 4V-aminopyrimidine ring forms strong hydrogen bonds between N1V and the side chain of E51 in the a domain and between N4Vand the carbonyl oxygen of G413 in the g domain. A weaker hydrogen bond exists between

Figure 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). (See color insert.)

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N3V and the nitrogen of I415 in the g domain, but the geometry is poor. The TDP is almost completely buried, so only the N4V exocyclic nitrogen of the 4V-aminopyrimidine ring and the C2 and S atoms of the thiazolium ring are accessible from the solvent. The TDP was found to be in the V-conformation (12), with the torsion angles deﬁning the relative orientations of the rings having values of ft =95.5j and fp = 65.9j. This conformation is never seen in small-molecule thiamine structures having an intact 4V-aminopyrimidine ring, except for thiamine thiazolone, which has a carbonyl at the thiazolium C2 position (13) that forms an internal hydrogen bond with N4V to provide stabilization. The V-conformation of the TDP is stabilized by several features of the protein environment including three hydrogen bonds to the 4V-aminopyrimidine ring (involving E51, G413, and I415) and also the presence of the bulky hydrophobic I415 side chain wedged beneath the two rings. These features of TDP binding were found conserved in all TDPdependent enzymes of known structure, although the type of hydrophobic residue beneath the rings varies. One important consequence of the V-conformation is to place the N4V and C2 atoms in very close proximity (3.4 A˚), which likely excludes the possibility that both N4V and C2 are completely protonated simultaneously. If this were the case, then the C2H and the nearest N4V hydrogen would be only 1.9 A˚ apart, well below the accepted van der Waals contact distance. The other apparently related role of the enzyme is to favor protonation at N1V in the 4V-aminopyrimidine ring. A strong hydrogen bond between the side chain of E51 and N1V stabilizes a tautomeric form of TDP in which the N4V amino group is converted to an imino group (1,2,14,15). The resulting eﬀect is to place an imino nitrogen atom in close proximity to the C2 hydrogen, with the basicity and geometry of the imino group ideal to abstract the proton at C2 and form the active ylide. Figure 7a shows steric crowding that would be occurring if the imino tautomer was not formed. Figure 7b shows the favorable interaction, with E51–N1V stabilizing the imino tautomeric form of the pyrimidine ring. The subsequent deprotonation of C2 is shown in Figure 7c. Although equilibrium almost certainly favors the state shown in Figure 7b, it was shown that the rate of proton transfer in the unfavored direction would be roughly 1 104 s1 M1. This still translates to a ﬁrstorder rate constant of more than 1 102 s1, which exceeds the turnover number; thus this deprotonation would not be rate limiting (2). No other suitable base is present in the active site to abstract the proton from C2, adding extra weight to the proposal that the V-conformation and the associated hydrogen bonding to the 4V-aminopyrimidine ring are crucial elements in the catalytic mechanism. The importance of the E51–N1V interaction has been conﬁrmed by studies in which substitutions were introduced at this point in YPDC (16–18) and at the corresponding location in transketolase (19).

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Figure 7 Potential ionization states of the TDP cofactor. (a) Isolated cofactor showing close contacts between hydrogen atoms present when in the 4V-amino form and in the V-conformation. (b) Tautomeric 4V-imino form of cofactor when enzyme bound and stabilized by interactions with E51 and G413. The hydrogen bond to G413 optimally orients N4V to point its lone pair at the C2 hydrogen. (c) In the 4V-amino form, stabilized by the enzyme and the fact that C2 is now deprotonated.

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III. SUBSTRATE ACTIVATION OF YEAST PYRUVATE DECARBOXYLASE A. The Substrate-Activation Pathway: Evidence in Solution As early as 1970, several groups had reported that YPDC is subject to substrate activation, as manifested in both the Hill coeﬃcient for v0–[S] plots and the lag phase in the pre-steady-state time course for NADH depletion (20,21). Schellenberger, Hu¨bner, and coworkers showed that there is a cysteine involved in substrate activation (22). With the help of a PDCI-6 fusion protein with only a single cysteine at position 221 (there are four cysteines in the wild-type enzyme at positions 69, 152, 221, and 222), it could be shown that this single cysteine was suﬃcient for substrate activation to exist (23). Next, the C221S, C222S singly, and C221S/C222S doubly substituted variants were created, and those with the C221S substitution were shown to no longer display substrate activation (24). These two experiments demonstrated that only residue C221 is both necessary and suﬃcient for substrate activation. Carrying out FT-IR and isoelectric focusing experiments in tandem, it was next shown that residue C221 is dissociated while residue H92 is protonated at pH 6.0, the optimum pH of YPDC (25). Despite the established need for C221 in YPDC activation, the mechanism by which allosteric eﬀects are mediated is unclear. The C221 residue is almost 20 A˚ away from the active site, so no direct interaction is possible. Rather, either binding information must be transferred via an interaction pathway or a signiﬁcant tertiary or quaternary structure change must take place. We have suggested and provided evidence for a potential pathway between C221 and the TDP that could cause modiﬁcation at C221 to have an allosteric aﬀect on the active site. Our proposed scheme is that the C221 nucleophile initially forms a hemithioketal adduct with the a-keto group of the substrate or activator. Upon binding, this adduct would interact unfavorably with the H92 side chain, causing either C221, H92, or both to move. Movement of H92 would then cause some displacement of E91. Residue E91 makes numerous contacts with a short segment of the g domain, including a hydrogen bond to the main-chain nitrogen of W412. This short segment consists of residues 410–415 and leads directly from the regulatory-site pocket at C221 to the active site. Residues 410–412 in this segment are in a position to interact with the regulatory pocket, whereas the top three, 413–415, are intimately related to TDP binding and form several of the key conserved protein–cofactor interactions. The proposed route of information transfer is shown in Figure 8. The importance of several of these residues in activation has been conﬁrmed by mutagenesis (23–30). The eﬀect of substitution at C221 was probed, suggesting that the C221S gave weakly negative cooperativity and the C221A or C221E(D) no

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Figure 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). (See color insert.)

cooperativity, while the wild-type C221 residue gives rise to positive cooperativity. Using solvent deuterium kinetic isotope eﬀect methods (see Chapter 13 in this volume), it was shown that the C221A substitution shifts the enzyme to an unactivated (lower activity) form (31), the C221D and C221E substitution shifts the enzyme to the activated form (32), and, very importantly, both unactivated and substrate-activated forms appear to have signiﬁcant activity, contrary to earlier claims (33). B. Is Quaternary Structure Related to the Activation Process? The proposed information transfer pathway from the regulatory site at C221 to the active site is present in both Forms A and B of the YPDC tetramer. This brings up the question of whether the change in quaternary structure has any relation to the proposed activation pathway. The ﬁrst observation of Form B of YPDC was seen in the presence of ketomalonate (6,7) and later in the presence of pyruvamide (3,8). This obviously brought forward the speculation that it was indeed the activators that caused the change in structure.

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However, more recently we have observed Form B in several other cases, including crystallization of YPDC variants, wild-type enzyme in the presence of inhibitors, and also wild-type enzyme in the absence of any eﬀectors at all! In addition we have, at times, found both A and B forms in the same crystallization drop for wild-type YPDC in the absence of any eﬀector molecules. Crystal data summarizing the occurrence of the two forms are shown in Table 1. Apparently, the energy barrier between the two forms is very small, since sometimes imperceptibly small changes in the enzyme or local solution environment can trigger form transitions. If quaternary structure is indeed related to activation, the question still remains as to how a signal at C221 could aﬀect quaternary structure so greatly. One possible explanation is that

Table 1

Crystal Data for YPDC Crystallized in the Presence and Absence of Eﬀectors

Source

Eﬀector/mutation

Sp gp

Z

Cell (A˚,j)

Form

Res (A˚)

81.0, 82.4, 116.6 69.5, 72.6, 62.4 141.9, 74.7, 119.9 90, 116.4, 90 142.0, 74.7, 120.0 90, 116.4, 90 80.3, 134.0, 117.3 90, 97.2, 90 92.0, 135.5, 99.3 90, 111.3, 90 145.1, 119.7, 81.3, 90, 120.2, 90 92.4, 136.3, 100.2 90, 111.4, 90

A

3.0

A

2.4

A

2.3

A

2.7

B

3.5

B

2.4

B

3.0

B

3.0

A

3.0

B

2.8

B

2.5

A

2.1

A

2.8

S. uvarum

None

P1

4

S. uvarum

None

C2

8

S. cerevisiae

None

C2

8

S. cerevisiae

P21

8

P21

8

C2

8

P21

8

S. cerevisiae

Ketomalonate (activator) Ketomalonate (activator) Pyruvamide (activator) p-chlorobenzylidene pyruvic acid (inhibitor) HgCl2 (inhibitor)

P21

8

S. cerevisiae

WT+C-termHis6

P1

4

S. cerevisiae

WT+C-termHis6

P21

8

S. cerevisiae

C221D,C222A

P21

8

S. cerevisiae

C221E,C222A

C2

8

S. cerevisiae

I415V

P1

4

S. cerevisiae S. cerevisiae S. cerevisiae

92.8, 135.9, 99.9 90, 111.6, 90 78.1, 80.7, 115.5 72.1, 70.0, 60.7 92.0, 133.8, 96.9 90, 111.3, 90 91.3, 133.7, 96.7 90, 111.6, 90 138.2, 75.1, 118.3 90, 118.2, 90 79.4, 79.3, 110.9 99.4, 103.6, 120.3

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the signal does not trigger the change in conformation per se but rather just stabilizes the readily accessible form B. Another possible explanation is that the signal at C221 does trigger the change in quaternary structure in the manner next described. Since the dimer–dimer interaction in Form A consists merely of a pair of hydrogen bonds on each side of the tetramer, it is possible that activation in some way acts to disrupt these bonds. The distance from C221 to the hydrogen bonds at the dimer interface is f18 A˚. However, C221 sits at the beginning of a short, poorly deﬁned 310 helical region of the protein that exhibits high thermal parameters. Residues D226 and E230 at the end of this poorly deﬁned region interact with K327, which lies at a position leading into of the last strand of the h domain, where the key dimer–dimer contacts are located in Form A. The potential signal pathway from C221 to the dimer– dimer interface at T320 is shown in Figure 9. Further, the thermal parameters of the 220–230 stretch are very diﬀerent between the two monomers in the crystallographic asymmetric unit in the original Form A structure (2), perhaps indicating that one of the monomers is indeed ready to undergo a structural rearrangement leading to a change in quaternary structure. In

Figure 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). (See color insert.)

Yeast Pyruvate Decarboxylase: New Features

187

addition, in the C221E/C222A crystal structure the electron density for this entire 220–230 region was very weak, indicating extreme ﬂexibility and/or disorder (to be published). To date we have been unable to obtain a covalent adduct at C221 that is visible in the electron density, and, contrary to our mutagenesis-derived evidence for the role of C221 in the allosteric mechanism, it was reported that when Form B crystallized in the presence of extremely high (300 mM) pyruvamide concentrations, the pyruvamide is not found as a covalent adduct at C221 but rather interacts with residues between the h and g domains almost 10 A˚ away (8). It should be pointed out, however, that in our attempts to crystallographically observe activators bound to C221, a reducing agent was always present in the media, which may have prevented stable binding from occurring. We also note that the mode of activator binding reported for the pyruvamide complex, although consistent with pyruvamide, would not be consistent with binding of the true substrate pyruvate. Since the amide donor nitrogen in pyruvamide would be replaced with a hydrogen bond accepting oxygen in pyruvate, the mode of binding could not be identical. Nevertheless, the binding of pyruvamide has been associated with the change in the relative orientation of the h domains and, more importantly, in ordering residues near the active site. Most signiﬁcantly, we must point out that the location of pyruvamide binding can aﬀect the same residues that are involved in both of our potential signaling pathways. Pyruvamide interacts directly with the R224 main-chain nitrogen, but an NH2 of the R224 side chain forms a strong hydrogen bond to the H92 carbonyl oxygen; thus the pyruvamide interaction could presumably trigger the same signal as proposed by a C221 adduct. Similarly, the pyruvamide interaction with R224 in the poorly deﬁned 310 helix outlined in the alternative activation pathway could again be potentially triggering that same pathway but by a diﬀerent mechanism. Studies with the C221A/C222A variant (31) suggested that pyruvamide interacts with YPDC in a diﬀerent manner than pyruvate, while other studies showed pyruvamide to be an inhibitor even at 80 mM concentration (32). Thus both of our postulated mechanisms of activation could be applicable to either C221 modiﬁcation by substrate or structural changes promoted by pyruvamide binding at its reported regulatory binding site. The exact nature of the substrate activation process is complicated and still largely unresolved, but it now seems highly likely that the changes in quaternary structure are indeed relevant to the activation process. C. Evidence for the Coexistence of Substrate-Activated and Active but Unregulated YPDC Conformations (34) YPDC exhibits a marked lag phase in the progress curves of product (acetaldehyde) formation. The earlier kinetic model by Schellenberger,

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Hu¨bner, and Schowen (SHS, ref. 35) for YPDC predicts that only upon binding of substrate in a regulatory site does a slow activation step convert inactive enzyme into the active form. This allosteric behavior gives rise to sigmoidal steady-state kinetics. The E477Q active-site variant of YPDC exhibited hyperbolic initial rate curves at low pH, not consistent with the model. Progress curves of product formation by this variant were S-shaped, consistent with the presence of three interconverting conformations with distinct steady-state rates. An example of the strong pH dependence of

Figure 10 Progress curves of acetaldehyde formation by WT YPDC at diﬀerent pH values. WT YPDC (10.5 Ag/mL) dissolved in pH-triple buﬀer, with diﬀerent pH values, containing 350 U/mL ADH, 0.3 mM NADH, 5 mM MgCl2, and 1 mM TDP was mixed in 1:1 ratio with a solution of 60 mM pyruvate.

Yeast Pyruvate Decarboxylase: New Features

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Scheme 2 Two-step phenomenological model of YPDC activation.

progress curves for acetaldehyde formation, one of the observations that led to our exploration of the new model, is shown in Figure 10. Surprisingly, wildtype YPDC at pH V 5.0 also possessed S-shaped progress curves, with the conformation corresponding to the middle steady state being the most active one. Re-examination of the activation by substrate of wild-type YPDC in the pH range of 4.5–6.5 revealed two characteristic transitions at all pH values. The phenomenological model for the two-step activation involving three conformational states of the enzyme, with two slow interconversions, is shown in Scheme 2. The values of steady-state rates are functions of both pH and substrate concentration, aﬀecting whether the progress curve appears ‘‘normal’’ or S-shaped with an inﬂection point. The substrate dependence of the apparent rate constants suggested that the ﬁrst transition corresponded to substrate binding in an active site and a subsequent step responsible for conversion to an asymmetric conformation. Consequently, the second enzyme state may report on ‘‘unregulated’’ enzyme, since the regulatory site does not participate in its generation. This enzyme state utilizes the alternating-sites mechanism, resulting in the hyperbolic substrate dependence of initial rate. Table 2 Kinetic Parameters of the Second Conformation (E1) for WildType YPDC pH 4.5 5.0 5.5 6.0 6.5

kcat (s1) 5.85 54.5 63.1 80.5 178.7

F F F F F

0.13 5.7 3.1 12.2 43.2

Km (mM)

kcat/Km (mM1 s1)

F F F F F

2.03 1.68 2.21 3.44 2.14

2.88 32.5 28.6 23.4 83.4

0.15 5.9 2.0 5.3 23.5

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The second transition corresponds to binding a substrate molecule in the regulatory site and subsequent minor conformational adjustments. The third enzyme state corresponds to the allosterically regulated conformation, previously referred to as activated enzyme. In Tables 2 and 3, it is shown that, depending on pH, the unregulated enzyme may have an activity exceeding that of even the substrate-activated form. The pH dependence of the Hill coeﬃcient suggests a random binding of pyruvate in a regulatory and an active site of wild-type YPDC. Addition of pyruvamide or acetaldehyde to YPDC results in the appearance of additional conformations of the enzyme. This new ﬁnding emphasizes the diﬃculties in deducing mechanistic information about this complex enzyme from kinetic studies. D. Further Support for Cysteine 221 as the Location of Substrate Activation on YPDC An interesting issue is why, in contrast to YPDC, the pyruvate decarboxylase from the bacterium Zymomonas mobilis (ZmPDC) and the closely related enzyme benzoylformate decarboxylase (BFD) are not substrate activated. The only direct protein chemical evidence we have for C221 was identiﬁcation of this site, where p-chlorocinnamaldehyde is covalently bound when derived from the mechanism-based inactivator p—ClC6H4CHjCHCOCOOH (28). In the X-ray structure of YPDC with 300 mM pyruvamide, the pyruvamide was found near the h domain (8) and at the active center, but the former site appears to be inappropriate on chemical grounds (see earlier section and Ref. 34). A comparison of the C221 region of YPDC with the corresponding regions of BFD and ZmPDC (34) provides a plausible explanation. While in YPDC, the putative tetrahedral thiohemiketal adduct between C221 and substrate forms an ion pair with H92 [reasonably mimicked by the C221D and C221E substitutions, which appear to shift the enzyme into a higher activity conformation according to kinetic criteria (32)]; on ZmPDC (61) and BFD

Table 3 Kinetic Parameters of the Third Conformation (E2) for Wild-Type YPDC pH 4.5 5.0 5.5 6.0 6.5 a

kcat (s1) 1.14 12.5 44.1 50.6 63.1

F F F F F

0.08 0.5 0.6 2.0 1.0

S0.5 (mM) 0.86 2.54 2.36 3.03 4.79

F F F F F

0.15 0.43 0.06 0.23 0.15

Parameter was ﬁxed to the value presented.

nH 0.72 F 1.00a 1.27 F 1.37 F 1.43 F

0.12 0.04 0.12 0.04

kcat/S0.5n

kcat/S0.5 (mM1 s1)

1.27 4.92 14.82 11.08 6.72

1.33 4.92 18.68 16.70 13.17

Yeast Pyruvate Decarboxylase: New Features

191

(62) the interactions are already present even in the absence of substrate. We have mapped the region of the mobile loop 291–303 (Wei et al., in preparation), and it indeed is crucial to maintaining activity, as already suggested in an earlier report (36). Hence, the h domain is important for maintaining activity of the enzyme, but substrate activation is only a modest, albeit subtle, superposition on this important role.

IV. EVIDENCE FOR A MECHANISM INVOLVING ALTERNATING ACTIVE SITES IN A FUNCTIONAL DIMER A. Development of the Model Our recent kinetic studies of YPDC variants substituted at positions D28 and E477 at the active center necessitate some modiﬁcation of the SHS (35) mechanism. It was found that enzyme without substrate activation apparently is still catalytically competent. Further, substrate-dependent inhibition of D28-substituted variants leads to an enzyme form with nonzero activity at full saturation, requiring a second major branch point in the mechanism. Kinetic data for the E477Q variant suggest that three consecutive substratebinding steps may be needed to release product acetaldehyde, unlikely if YPDC monomer with a single catalytic and regulatory site for substrate binding is the minimal catalytic unit. A model to account for all kinetic observations involves a functional dimer operating through alternation of active sites. In this mechanism, roles were suggested for the active-center acid– base groups D28, E477, H114, and H115 (37). The appearance of several substrate-binding sites in steady-state kinetics requires that, in addition to the mere presence of those binding sites, they inﬂuence each other’s binding or catalytic properties. The pathway of signal transduction from the regulatory site to the active site of YPDC deduced in our laboratories was summarized earlier; most likely, it links two sites of the same subunit. The presence of the third binding site for pyruvate implies the existence of an additional signal transduction pathway connecting two diﬀerent subunits. The two active-site acid–base residues D28 and E477 appear to be important both for catalysis and for the postulated signal transduction pathway. Variants of both D28 and E477 residues gave evidence of the presence of elevated concentration of postdecarboxylation intermediates, suggesting that the accumulation of those intermediates, and the unusual kinetic behavior of those variants, is the result of both a catalytic and a regulatory role of those residues. The new mechanism is based on the principles exploited in an internal combustion engine, suggested for enzymes, which sustain some functional

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locomotion (38,39); we also utilized the same type of mechanism to explain the pre-steady-state kinetic behavior of BFD (40,41). Substrate binding in the second active site of BFD was required for the decarboxylation of the benzoylformic acid–TDP covalent intermediate in the ﬁrst binding site. The same model could be utilized for the active sites of YPDC: Decarboxylation in one active site may supply the energy for a second active site to bind substrate or to release product, making the sequential operation a prerequisite for normal function. Two active sites of the functional homodimer of YPDC have diﬀerent conformations in the pyruvamide-complexed structure (8), so it is possible that the diﬀerence in the geometry of the active sites is a reﬂection of their transiently diﬀerent function during catalysis. Since D28 is within hydrogen-bonding distance from H114 or H115, its ionization state may change as a consequence of changing the distance from those residues. It is also possible that D28-COOH, being close to the covalently bound substate-TDP or product-TDP adduct, may lose or gain a proton as part of the catalytic cycle. We concluded in the study of carboligase side reactions (42) that D28 indeed changes its ionization state from D28-COOH to D28-COO once the decarboxylation is completed, ensuring that the charge of the D28 residue reﬂects the functional state of the active site. We tentatively concluded that D28 and E477Q might be in the signal transduction pathway, which keeps the work of alternating sites in sequence. The following mechanism of alternating sites was formulated. Let us suppose that both active sites catalyze the full catalytic cycle, with a mandatory phase shift between two active sites of the functional dimer: One active site of the functional dimer catalyzes the reactions steps through decarboxylation, the second subunit at the same instant catalyzes the postdecarboxylation phase of the reaction. Further, the coupling of the two phases of the reactions is mediated through both the prerequisite change in the conformation and the charge/hydrogen bond pattern of the active site, as well as thermodynamic requirements of the various steps of the reaction. Irreversibility of the decarboxylation reaction not only makes it thermodynamically favorable but also provides a possible source of energy for the endergonic parts of the reaction or for the shift of an unfavorable equilibrium to enhance the overall throughput of the enzyme. Alternation of sites through a conformational change would ensure the physical coupling of the two parts of the reaction. As seen in Section II, there is evidence that multiple conformations are readily accessible to YPDC, with little energetic penalty. While we cannot assign the two speciﬁc reaction steps that are coupled, we believe that each of them represents two phases of the overall reaction, namely, predecarboxylation and postdecarboxylation, as illustrated in Scheme 3. In this scheme, E represents activated enzyme, i.e., ES* in the SHS formalism, where S is free substrate. ES and EN represent one of the

Yeast Pyruvate Decarboxylase: New Features

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Scheme 3 A model of the alternating-sites mechanism.

predecarboxylation (enzyme–substrate noncovalent Michaelis complex or LTDP) and postdecarboxylation (enamine or HETDP, and the enzyme– product noncovalent complex) intermediates, respectively. The positioning of the letter S or N to the right or left side of the symbol for the enzyme reﬂects involvement of two diﬀerent monomers of the functional dimer. As can be seen, the release of product is possible only after substrate binding and, possibly, LTDP formation in the neighboring subunit. Some of the enzyme species (for example, ES and SEN) represent nodes (vertices) of the graph wih the choice of reactions they can undergo, and they are partitioned between diﬀerent pathways according to their relative kinetic eﬃcacy. The intermediate EN is formed in an irreversible step from decarboxylation of ES. The product release is irreversible due to reduction of acetaldehyde (P) in the coupled reaction. If either of the two irreversible steps is blocked, an accumulation of pre- or postdecarboxylation intermediates will result. Notice that, for simplicity, Scheme 3 does not include catalysis by nonactivated enzyme, although this would be a logical extension of the model (as seen earlier, evidence from this laboratory suggests that this enzyme form is indeed active). We suggest that the pH eﬀects on the steady-state kinetics of the wildtype YPDC and its variants highlight diﬀerent parts of a single mechanistic model due to changes in relative kinetic eﬃciency of particular steps. The proposed mechanistic model of the reaction represents the network of possible reversible and irreversible steps. This same model could explain the observed hyperbolic behavior as well as the enhanced cooperativity with a

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Hill coeﬃcient greater than 2. It should be noted that binding of pyruvate in the regulatory site of the second monomer of the dimer is not included in Scheme 3. Therefore, each catalytic cycle may include an additional substratebinding step for saturation of the regulatory site, a consequence being a Hill coeﬃcient as high as 4, providing that particular conditions will favor substrate-binding steps and LTDP formation of the activated enzyme over decarboxylation. Elsewhere, we presented kinetic expressions to the several scenarios outlined in Scheme 3 and discussed how the various scenarios could account for our observations with the D28A and E477Q variants as well as those for the wild-type enzyme (37). The observations with pyruvamide can also be explained in light of the alternating-sites model. In addition to binding in the regulatory site (Ka), pyruvamide may also bind in the second active site of the enzyme species ES (step ks5). This will lead to the lower catalytic turnover rate, as does inhibition by pyruvate. Pyruvamide could inhibit both wild-type and variant YPDCs by mimicking pyruvate, by forming a PA.ES complex, analogous to the SES complex in Scheme 3. B. Activation in the Alternating-Active-Sites Model The hypothesis of alternating-catalytic-sites reactivity is buttressed by the close contact of the subunits in the asymmetrical dimer (8; PDB accession 1QPB), and the striking diﬀerences between the two monomers. We suggest that activation of YPDC is a necessary event to start the catalytic cycle, though it may not be required for its perpetuation. Certainly, the activation step cannot be a part of the catalytic cycle of the E477Q and D28N variants, whose carboligase reaction was studied in detail (42). It was reported that the rate constant for the zero-order activation reaction of wild-type YPDC (subsequent to substrate binding in the regulatory site) is 0.5 s1. Should this step be an essential part of each catalytic cycle, the turnover could not be higher than this value. For the E477Q and D28N variants the turnover number for acetoin formation is twice this number. One can certainly expect that substitution of diﬀerent amino acid residues may result in a change of the rate of the activation process. However, it is diﬃcult to explain why substitutions at two diﬀerent active-site positions would lead to the same magnitude of rate enhancements. In addition, only one regulatory site is clearly visible in the steady-state kinetic behavior of the variants. Therefore, we are compelled to suggest that the activation step, which changes the enzyme from a lower to a higher activity form, is required to commence the catalytic cycle, i.e., E represents the higher-activity conformation. But the higher-activity conformation persists through the catalytic

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cycle, whether or not pyruvate remains in the regulatory binding site. The aﬃnity of the regulatory site for the substrate/activator is likely to increase as a result of the occupation of the active site. C. Structural Hints Suggesting Alternating Active-Sites Reactivity Mechanistically, formation of LTDP from pyruvate and TDP and the release of acetaldehyde from HETDP represent, in essence, the same reaction in opposite directions, the transfer of a proton to or from the C2a oxygen atom. These reactions are preceded by proton abstraction from C2 of the thiazolium ring and are separated by proton donation to the C2a atom of the enamine to form HETDP. The latter two reactions also represent the reverse of mechanistically very similar reactions. Therefore, the sequence of reactions carried out by YPDC can be viewed as two reactions in tandem, where in the second repeat the reverse reactions are utilized. The four parts of the catalytic cycle are shown in Scheme 4, where parts 1 and 3 represent reversal of one reaction and parts 2 and 4 represent reversal of another reaction. Invoking the principle of microscopic reversibility, it would be more eﬃcient for the enzyme to utilize the same group for each pair of the reversed reaction. Although, a priori, utilization of the same group in all four reactions is not a requirement, an inspection of the YPDC structure suggests that this is an attractive possibility.

Scheme 4

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Activation perturbs YPDC from its nearly symmetrical state to a highly asymmetrical one. In each dimer of the 1QPB structure of YPDC crystallized with 300 mM pyruvamide (8), one active site reveals a closed, the other an open conformation. On the basis of the structural data, we propose a model where this same N4V-amino/imino (see Sec. IV for evidence) group of TDP participates in all of the reactions. The choice of the candidate for this reaction should reﬂect easy regulation of pKa of the group. Since only E477 and D28 of the four potential acid–base side chains are in close proximity to the C2 of the thiazolium ring, we select these, rather than H114 or H115. We assume that the N4V atom of the 4V-aminopyrimidine ring is the major player in the proton transfer in the ﬁrst half of the reaction. With the V-conformation enforced by several factors (43), it is virtually impossible to displace the amino group by several angstroms. The ‘‘closed’’ active site of the A/C subunit conformation can accommodate proton transfer from C2 to N4V, but the site is too tight to accommodate the covalently bound pyruvate of LTDP. Therefore, we suggest that the A/C conformation of the closed active site provides a working model for formation of both the ylide and HETDP, where the N4V atom comes within close contact of the C2 or C2a atom to donate or accept a proton. In the ‘‘open’’ active site of the B/O subunit conformation, the N4V atom is further from C2a and provides a model for participation of N4V in the protonation or deprotonation of the C2a-hydroxyl group in LTDP formation or acetaldehyde release. According to this working hypothesis, D28 and E477 (as well as H114 and H115) will assist in the proton transfer, providing a hydrogen relay chain. Neither the E477Q nor the D28A substitutions could change the rate of C2H/D exchange in the nonactivated enzyme (44). However, the rate of exchange in the presence of pyruvamide was 20-fold lower for those variants than for the wild-type YPDC (45), suggesting that the reaction pathways may not be identical in the activated and nonactivated enzymes. At the same time, these results also hinted at the participation in some manner of both D28 and E477 in the signal transduction pathway, leading from the regulatory, or perhaps inhibitory, site for pyruvamide to the active site. In the ‘‘closed’’ A/C active site, the nearest distance between D28 and N4V is 6.65 A˚ and between D28 and E477 is 5.85 A˚. In the B/O (open) subunit, the minimal distance from D28 to N4V is 4.98 A˚ and from D28 to E477 is 3.84 A˚, suggesting that the rate of proton transfer between N4V and D28 and/or E477 may be enhanced in the B/O subunit. It would be important, for example, to transfer the proton to the N4V atom once LTDP was formed, to prevent the loss of pyruvate leading to reversal of the reaction. This may also be a convenient mechanism for changing the state of ionization of the D28 side chain if D28COOH were to donate the proton. The decarboxylation of

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LTDP to the enamine follows, rendering the enamine ready to accept a proton from the N4V atom, and D28–COO will participate in the protection of postdecarboxylation intermediates (42). Invoking the principle of microscopic reversibility, if the N4V-amino group is involved in protonation of the C2a-alkoxide of LTDP, it might also participate in the release of acetaldehyde, abstracting a proton from the C2ahydroxyl group of HETDP. However, it is likely that one of the residues among D28, H114, H115, and E477 needs to assist in the latter reaction; acetaldehyde release was impaired to a greater degree by the alteration of these active-site residues than the steps preceding decarboxylation, judging by the rates of acetoin formation. Just as was reasoned for LTDP formation, it would be important, for example, to transfer a proton from the N4V atom, once acetaldehyde is released, so as to prevent the reversal with HETDP formation. Once again, the D28 side chain will become D28-COOH, signaling the end of the postdecarboxylation phase. And the imino-tautomer of the 4Vaminopyrimidine ring regenerated in this reaction will be ready for catalysis of the ﬁrst step of the catalytic cycle. It appears that H114 and/or H115 may create the site for the inhibitory pyruvate molecule (inhibition evident from the steady-state kinetics), for the following reason. We deduced that prior to decarboxylation, the charge distribution at the active site is H115+D280, which is useful for attracting the ﬁrst pyruvate and to form the LTDP. Once decarboxylation is completed, the charge distribution was deduced to change to H115+ D28, so as to repel the second pyruvate that could form acetolactate in an undesirable side reaction. With the D28A and D28N variants, the charge distribution would be the same during the entire reaction sequence, H115+D280, which could indeed be more attractive for a pyruvate molecule in either active site. The Hill coeﬃcient of both E477Q and H115F variants displayed similar pH dependencies (44). Given that both E477 and H115 likely form hydrogen bonds to the residue D28, this similarity of Hill coeﬃcient behavior may reﬂect that participation of both amino acids in the interaction between active sites is expressed through the residue D28. Therefore, it is possible that the three acid–base residues (D28, H115, E477, and perhaps H114) of the active site participate in signal transduction from one active site to another. An interesting, still unresolved mechanistic issue concerns the step in the mechanism where hydroxide ion is released to the solution. The sum of the evidence suggested that protonation of the ylide by water, once acetaldehyde is released, could be a likely candidate, since the ylide is certainly a strong base. Were this correct, ylide formation, present as a part of the cycle, might be the rate-limiting step under some conditions. The two variants E477Q and D28A and wild-type YPDC all gave the same rate of C2H/D exchange

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in nonactivated enzyme: 0.9–1.0 s1 at 5jC or approximately 10 s1 at room temperature (44). The C221A and C221S variants are suspected to be nonactivated during the entire catalytic cycle. Their activity is about 25% of the wild-type enzyme; i.e., it is possible that ylide formation in nonactivated enzyme is the rate-limiting step (31). At the same time, C2H/D exchange in activated YPDC was reported to be > 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

REFERENCES 1.

2.

F Dyda, W Furey, S Swaminathan, M Sax, B Farrenkopf, F Jordan. Catalytic centers in the thiamin diphosphate–dependent enzyme pyruvate decarboxylase at 2.4-A˚ resolution. Biochemistry 32:6165–6170, 1993. 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

Yeast Pyruvate Decarboxylase: New Features

3.

4.

5. 6.

7.

8.

9. 10.

11. 12.

13.

14.

15.

16.

211

diphosphate–dependent enzyme pyruvate decarboxylase from the yeast Saccharomyces cerevisiae at 2.3-A˚ resolution. J Mol Biol 256:590–600, 1996. 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. 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 issue 2, in press, 2003. W Furey, P Arjunan, L Chen, F Dyda, T Umland, S Swaminathan, M Sax, F Jordan, B Farrenkopf, Y Gao, D Zhang. Multiple modes of tetramer assembly and insight into allosteric activation revealed by X-ray crystal structures of pyruvate decarboxylase. In: H Bisswanger, A Schellenberger eds. Biochemistry and Physiology of Thiamin Diphoshapte Enzymes. Prien, Germany: A. u. C. Intermann Wissenschaftlicher Verlag, pp. 103–124, 1996. W Furey, P Arjunan, L Chen, M Sax, F Guo, F Jordan. Structure–function relationships and ﬂexible tetramer assembly in pyruvate decarboxylase revealed by analysis of crystal structures. Biochim Biophys Acta 1385:253–270, 1998. 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. CF Hawkins, A Borges, RN Perham. A common structural motif in thiamin pyrophosphate–binding enzymes. FEBS Lett 255:77–82, 1989. Y Lindqvist, G Schneider, U Ermler, M Sundstrom. Three-dimensional structure of transketolase, a thiamine diphosphate–dependent enzyme, at 2.5-A˚ resolution. Embo J 11:2373–2379, 1992. YA Muller, GE Schulz. Structure of the thiamine- and ﬂavin-dependent enzyme pyruvate oxidase. Science 259:965–967, 1993. J Pletcher, M Sax, G Blank, M Wood. Stereochemistry of intermediates in thiamine catalysis. 2. Crystal structure of DL-2-(a-hydroxybenzyl)thiamine chloride hydrochloride trihydrate. J Am Chem Soc 99:1396–1403, 1977. W Shin, YC Kim. Crystal structure of thiamin thiazolone: a possible transitionstate analogue with an intramolecular N—H. . .O hydrogen bond in the V form. J Am Chem Soc 108:7078–7082, 1986. 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. 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. F Jordan, I Baburina, Y Gao, A Kahyaoglu, N Nemeria, A Volkov, J Yi, D Zhang, R Machado, J Guest, W Furey, S Hohmann. New insights to the regulation of thiamin diphosphate dependent decarboxylases by substrate and ThDP.Mg(II). In: H Bisswanger, A Schellenberger, eds. Biochemistry and Phys-

212

17.

18.

19.

20. 21.

22.

23.

24.

25.

26.

27.

28.

Jordan et al. iology of Thiamin Diphosphate Enzymes. 1996. Prien, Germany: A. u. C. Intermann Wissenschaftlicher Verlag, pp. 53–69, 1996. M Killenberg-Jabs, S Ko¨nig, I Eberhardt, S Hohmann, G Hu¨bner. Role of Glu51 for binding and catalytic activity in pyruvate decarboxylase from yeast studied by site-directed mutagenesis. Biochemistry 36:1900–1905, 1997. Y Gao. Mechanism of Pyruvate Decarboxylase-Site-Directed Mutagenesis Studies on Glu51. PhD dissertation, Rutgers University Graduate Faculty at Newark, NJ, 2000. C Wikner, L Meshalkina, U Nilsson, M Nikkola, Y Lindqvist, M Sundstrom, G Schneider. Analysis of an invariant cofactor–protein interaction in thiamin diphosphate–dependent enzymes by site-directed mutagenesis. Glutamic acid 418 in transketolase is essential for catalysis. J Biol Chem 269:32144–32150, 1994. A Boiteux, B Hess. Allosteric properties of yeast pyruvate decarboxylase. FEBS Lett 9:293–296, 1970. J Ullrich, I Donner. Kinetic evidence for two active sites in cytoplasmic yeast pyruvate decarboxylase. Hoppe Seylers Z Physiol Chem 351:1026–1029, 1970. 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. X Zeng, B Farrenkopf, S Hohmann, F Dyda, W Furey, F Jordan. Role of cysteines in the activation and inactivation of brewer’s yeast pyruvate decarboxylase investigated with a PDC1-PDC6 fusion protein. Biochemistry 32:2704– 2709, 1993. 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 221 to serine. Biochemistry 33:5630–5635, 1994. I Baburina, DJ Moore, A Volkov, A Kahyaoglu, F Jordan, R Mendselsohn. 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. F Jordan, N Nemeria, F Guo, I Baburina, Y Gao, A Kahyaoglu, H Li, J Wang, J Yi, 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. I Baburina, H Li, B Bennion, W Furey, F Jordan. Interdomain information transfer during substrate activation of yeast pyruvate decarboxylase: the interaction between cysteine 221 and histidine 92. Biochemistry 37:1235–1244, 1998. I Baburina, G Dikdan, F Guo, G Tous, B Root, F Jordan. Reactivity at the substrate activation site of yeast pyruvate decarboxylase: inhibition by distortion of domain interactions. Biochemistry 37:1245–1255, 1998.

Yeast Pyruvate Decarboxylase: New Features

213

29. H Li, W Furey, F Jordan. Role of glutamate 91 in information transfer during substrate activation of yeast pyruvate decarboxylase. Biochemistry 38:9992– 10003, 1999. 30. H Li, F Jordan. Eﬀects of substitution of tryptophan 412 in the substrate activation pathway of yeast pyruvate decarboxylase. Biochemistry 38:10004– 10012, 1999. 31. 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. 32. 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. 33. 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. 34. EA Sergienko, F Jordan. A new model for activation of yeast pyruvate decarboxylase by substrate consistent with the alternating-sites mechanism: demonstration of the existence of two active forms of the enzyme. Biochemistry 41:3952–3967, 2002. 35. 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 113:8402–8409, 1991. 36. I Eberhardt, H Cederberg, S Ko¨nig, H Li, F Jordan, S Hohmann. Autoregulation of yeast pyruvate decarboxylase gene expression requires the enzyme but not its catalytic activity. Eur J Biochem 262:191–202, 1999. 37. 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. 38. PD Boyer. The ATP synthase—a splendid molecular machine. Annu Rev Biochem 66:717–749, 1997. 39. KM Brendza, DJ Rose, SP Gilbert, WM Sexton. Lethal kinesin mutations reveal amino acids important for ATPase activation and structural coupling. J Biol Chem 274:31506–31514, 1999. 40. EA Sergienko, J Wang, L Polovnikova, MS Hasson, MJ McLeish, GL Kenyon, F Jordan. Spectroscopic detection of transient thiamin diphosphate– bound inter-mediates on benzoylformate decarboxylase. Biochemistry 39: 13862–13869, 2000. 41. L Polovnikova, MJ McLeish, EA Sergienko, JT Burgner, NL Anderson, AK Bera, F Jordan, GL Kenyon, MS Hasson. Structural and kinetic analysis of catalysis by a thiamin diphosphate–dependent enzyme, benzoylformate decarboxylase. Biochemistry 42:1820–1830, 2003. 42. EA Sergienko, F Jordan. Catalytic acid–base groups in yeast pyruvate de-

214

43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

Jordan et al. carboxylase 2. Insights to the speciﬁc roles of D28 and E477 from the rates and stereospeciﬁcity of formation of carboligase side products. Biochemistry 40: 7369–7381, 2001. F Guo, D Zhang, A Kahyaoglou, RS Farid, F Jordan. Is a hydrophobic amino acid required to maintain the reactive V-conformation 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. M Liu, EA Sergienko, F Guo, J Wang, K Tittmann, G Hu¨bner, 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. D Kern, G Kern, H Neef, K Tittmann, M Killenberg-Jabs, C Wikner, G Schneider, G Hu¨bner. How thiamine diphosphate is activated in enzymes. Science 275:67–70, 1997. J Yi, N Nemeria, A McNally, F Jordan, RS Machado, JR Guest. Eﬀect of substitutions in the thiamin diphosphate–magnesium fold on the activation of the pyruvate dehydrogenase complex from Escherichia coli by cofactors and substrate. J Biol Chem 271:33192–33200, 1996. N Nemeria, A Volkov, A Brown, J Yi, L Zipper, JR Guest, F Jordan. Systematic study of all six cysteines of the E1 subunit of the pyruvate dehydrogenase multienzyme complex from Escherichia coli: none is essential for activity. Biochemistry 37:911–922, 1998. EA Sergienko, F Jordan. Yeast pyruvate decarboxylase tetramers can dissociate into dimers along two interfaces. Hybrids of low-activity D28A (or D28N) and E477Q variants, with substitution of adjacent active-center acidic groups from diﬀerent subunits, display restored activity. Biochemistry 41:6164– 6169, 2002. S Zhang. Studies on the Key Thiamin Diphosphate-bound Enamine Intermediate in Models and Enzymes. PhD dissertation, Rutgers University Graduate Faculty at Newark, NJ 2003. F Jordan, G Chen, S Nishikawa, B Sundoro-Wu. Potential roles of the aminopyrimidine ring in thiamin-catalyzed reactions. Ann New York Acad Sci 378:14–31, 1982. F Jordan. 1H NMR Evidence for high barriers to amino group rotation in 4Vaminopyrimidines, including thiamin, at low pH in water. J Org Chem 47:2748– 275, 1982. MV Kovina, IA Bykova, ON Solovjeva, LE Meshalkina, GA Kochetov. The origin of the absorption band induced through the interaction between apotransketolase and thiamin diphosphate. Biochem Biophys Res Commun 294:155–160, 2002. 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.

Yeast Pyruvate Decarboxylase: New Features

215

54. K Tittmann, R Golbik, K Uhlemann, L Khailova, G Schneider, MS Patel, F Jordan, D Chipman, RG Duggleby, G Hu¨bner. Snapshots of thiamindependent enzymes in action (submitted); and K Tittmann et al., Chapter 5, this volume. 55. M Liu. Studies on Yeast Pyruvate Decarboxylase. Function of the active Center Acid-Base Groups. Partitioning of Thiamin-bound Covalent Intermediates. PhD dissertation, Rutgers University Graduate Faculty at Newark, NJ, 2002. 56. Z Zhang. Studies on Yeast Pyruvate Decarboxylase. Intermediate Partitioning. A Model for the 1V,4V-Iminothiamin Diphosphate Tautomer. PhD dissertation, Rutgers University Graduate Faculty at Newark, NJ, 2002. 57. G Barletta, WP Huskey, F Jordan. Observation of a 2-a-enamine from a 2[a-methoxy-a-phenylmethyl]-3,4-dimethylthiazolium salt in water: implications for catalysis by thiamin diphosphate–dependent a-keto acid decarboxylases. J Am Chem Soc 114:7607–7608, 1992. 58. G Barletta, WP Huskey, F Jordan. Ionization kinetics at the C2a position of 2benzylthiazolium salts leading to examines relevant to thiamin-catalyzed enzymatic reactions. J Am Chem Soc 119:2356–2362, 1997. 59. Y Zou. Investigating the Mechanism of Catalysis in Thiamin-dependent Enzymes. PhD dissertation, Rutgers University Graduate Faculty at Newark, NJ, 1999. 60. F Jordan, H Li, A Brown. Remarkable stabilization of zwitterionic intermediates may account for a billion-fold rate acceleration by thiamin diphosphate– dependent decarboxylases. Biochemistry 38:6369–6373, 1999. 61. 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. 62. 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. 63. M Carson. RIBBONS2.0. J Appl Crystallogr 24:958–961, 1991.

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) at 0.5/1 = 0.5. Alternatively, one example of a case giving a normal (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.

Isotope Effects on Yeast Pyruvate Decarboxylase 6.

7.

8.

9. 10.

11. 12. 13.

14. 15. 16.

17.

18.

19.

20.

21.

229

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.

230

Wei et al.

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.

Isotope Effects on Yeast Pyruvate Decarboxylase

231

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.

3.

4.

5.

6.

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.

Mechanism and Regulation of Bacterial AHASs 7.

8.

9.

10.

11. 12.

13. 14.

15.

16.

17. 18. 19. 20. 21.

22.

23.

249

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.

250

Chipman et al.

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.

4.

RG Duggleby, SS Pang. Acetohydroxyacid synthase. J Biochem Molec Biol 33:1–36, 2000. FC Sto¨rmer. The pH 6 acetolactate-forming enzyme from Aerobacter aerogenes. II. Evidence that it is not a ﬂavoprotein. J Biol Chem 243:3740–3741, 1968. A Joutel, A Ducros, S Alamowitch, C Cruaud, V Domenga, E Mare´chal, K Vahedi, H Chabriat, MG Bousser, E Tournier-Lasserve. A human homolog of bacterial acetolactate synthase genes maps within the CADASIL critical region. Genomics 38:192–198, 1996. RG Duggleby, AER Kartikasari, RM Wunsch, YT Lee, MW Kil, JY Shin, SI Chang. Expression in Escherichia coli of a putative human acetohydroxyacid synthase. J Biochem Molec Biol 33:195–201, 2000.

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6.

7.

8. 9.

10.

11.

12. 13.

14.

15.

16. 17.

18. 19.

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V Foulon, VD Antonenkov, K Croes, E Waelkens, GP Mannaerts, PP Van Veldhoven, M Casteels. Puriﬁcation, molecular cloning, and expression of 2-hydroxyphytanoyl-CoA lyase, a peroxisomal thiamine pyrophosphate-dependent enzyme that catalyzes the carbon–carbon bond cleavage during alphaoxidation of 3-methyl-branched fatty acids. Proc Natl Acad Sci USA 96:10039– 10044, 1999. JA Grandoni, PT Marta, JV Schloss. Inhibitors of branched-chain amino acid biosynthesis as potential antituberculosis agents. J Antimicrob Chemother 42:475–482, 1998. RA LaRossa, JV Schloss. The sulfonylurea herbicide sulfometuron methyl is an extremely potent and selective inhibitor of acetolactate synthase in Salmonella typhimurium. J Biol Chem 259:8753–8757, 1984. DL Shaner, PC Anderson, MA Stidham. Imidazolinones: potent inhibitors of acetohydroxyacid synthase. Plant Physiol 76:545–546, 1984. JK Smith, JV Schloss, BJ Mazur. Functional expression of plant acetolactate synthase genes in Escherichia coli. Proc Natl Acad Sci USA 86:4179–4183, 1989. P Bernasconi, AR Woodworth, BA Rosen, MV Subramanian, DL Siehl. A naturally occurring point mutation confers broad range tolerance to herbicides that target acetolactate synthase. J Biol Chem 270:17381–17385, 1995. MW Kil, SI Chang. Expression in Escherichia coli, puriﬁcation, and characterization of the tobacco sulfonylurea herbicide-resistant recombinant acetolactate synthase and its interaction with the triazolopyrimidine herbicides. J Biochem Molec Biol 31:287–293, 1998. AK Chang, RG Duggleby. Expression, puriﬁcation and characterization of Arabidopsis thaliana acetohydroxyacid synthase. Biochem J 327:161–169, 1997. CM Hill, SS Pang, RG Duggleby. Puriﬁcation of Escherichia coli acetohydroxyacid synthase isozyme II and reconstitution of active enzyme from its individual pure subunits. Biochem J 327:891–898, 1997. SS Pang, RG Duggleby. Expression, puriﬁcation, characterization and reconstitution of the large and small subunits of yeast acetohydroxyacid synthase. Biochemistry 38:5222–5231, 1999. L Eoyang, PM Silverman. Puriﬁcation and subunit composition of acetohydroxyacid synthase I from Escherichia coli K-12. J Bacteriol 157:184–189, 1984. 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. RG Duggleby. Identiﬁcation of an acetolactate synthase small subunit gene in two eucaryotes. Gene 190:245–249, 1997. B Singh, I Szamosi, JM Hand, R Misra. Arabidopsis acetohydroxyacid synthase expressed in Escherichia coli is insensitive to the feedback inhibitors. Plant Physiol 99:812–816, 1992.

272

Duggleby et al.

20. C Poulsen, P Stougaard. Puriﬁcation and properties of Saccharomyces cerevisiae acetolactate synthase from recombinant Escherichia coli. Eur J Biochem 185:433–439, 1989. 21. AG Hinnebusch. The general control of amino acid biosynthetic genes in the yeast Saccharomyces cerevisiae. CRC Crit Rev Biochem 21:277–317, 1986. 22. C Cullin, A Baudin-Baillieu, E Guillemet, O Ozier-Kalogeropoulos. Functional analysis of YCL09c: evidence for a role as the regulatory subunit of acetolactate synthase. Yeast 12:1511–1518, 1996. 23. PT Magee, H de Robichon-Szulmajster. The regulation of isoleucine-valine biosynthesis in Saccharomyces cerevisiae. 3. Properties and regulation of the activity of acetohydroxyacid synthetase. Eur J Biochem 3:507–511, 1968. 24. DF Caroline, RW Harding, H Kuwana, T Satyanarayana, RP Wagner. The iv3 mutants of Neurospora crassa. II. Activity of acetohydroxy acid synthetase. Genetics 62:487–494, 1969. 25. L Glatzer, E Eakin, RP Wagner. Acetohydroxy acid synthetase with a pH optimum of 7.5 from Neurospora crassa mitochondria: characterization and partial puriﬁcation. J Bacteriol 112:453–464, 1972. 26. S Takenaka, H Kuwana. Control of acetohydroxy acid synthetase in Neurospora crassa. J Biochem 72:1139–1145, 1972. 27. H Kuwana, M Date. Solubilization of valine-sensitive acetohydroxy acid synthetase from Neurospora mitochondria. J Biochem 77:257–259, 1975. 28. SS Pang, RG Duggleby. Regulation of yeast acetohydroxyacid synthase by valine and ATP. Biochem J 357:749–757, 2001. 29. HP Hershey, LJ Schwartz, JP Gale, LM Abell. Cloning and functional expression of the small subunit of acetolactate synthase from Nicotiana plumbaginifolia. Plant Molec Biol 40:795–806, 1999. 30. YT Lee, RG Duggleby. Identiﬁcation of the regulatory subunit of Arabidopsis thaliana acetohydroxyacid synthase and reconstitution with its catalytic subunit. Biochemistry 40:6836–6844, 2001. 31. Y Oda, Y Nakano, S Kitaoka. Properties and regulation of valine-sensitive acetolactate synthase from mitochondria of Euglena gracilis. J Gen Microbiol 128:1211–1216, 1982. 32. FE Dailey, JE Cronan Jr. Acetohydroxy acid synthase I, a required enzyme for isoleucine and valine biosynthesis in Escherichia coli K-12 during growth on acetate as the sole carbon source. J Bacteriol 165:453–460, 1986. 33. 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. 34. RP Lawther, RC Wek, JM Lopes, R Pereira, BE Taillon, GW Hatﬁeld. The complete nucleotide sequence of the ilvGMEDA operon of Escherichia coli K12. Nucleic Acids Res 15:2137–2155, 1987. 35. P Meuris. Studies of mutants inhibited by their own metabolites in Saccharomyces cerevisiae II. Genetic and enzymatic analysis of three classes of mutants. Genetics 63:569–580, 1969. 36. C Larsson, A Nilsson, A Blomberg, L Gustafsson. Glycolytic ﬂux is

Structure and Properties of AHAS

37.

38.

39. 40.

41.

42.

43.

44.

45.

46.

47.

48. 49. 50.

51.

273

conditionally correlated with ATP concentration in Saccharomyces cerevisiae: a chemostat study under carbon- or nitrogen-limiting conditions. J Bacteriol 179:7243–7250, 1997. SS Pang, LW Guddat, RG Duggleby. Crystallization of the catalytic subunit of Saccharomyces cerevisiae acetohydroxyacid synthase. Acta Cryst D57:1321– 1323, 2001. SS Pang, RG Duggleby, LW Guddat. Crystal structure of yeast acetohydroxyacid synthase: a target for herbicidal inhibitors. J Molec Biol 317:249–262, 2002. YT Lee, RG Duggleby. Regulatory interactions in Arabidopsis thaliana acetohydroxyacid synthase. FEBS Lett 512:180–184, 2002. 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. YY 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 Molec Biol 237:315–335, 1994. SC Falco, RE McDevitt, CF Chui, ME Hartnett, S Knowlton, CJ Mauvais, JK Smith, BJ Mazur. Engineering herbicide-resistant acetolactate synthase. Dev Ind Microbiol 30:187–194, 1989. AK Chang, RG Duggleby. Herbicide-resistant forms of Arabidopsis thaliana acetohydroxyacid synthase: characterization of the catalytic properties and sensitivity to inhibitors of four deﬁned mutants. Biochem J 333:765–777, 1998. CM Hill, RG Duggleby. Mutagenesis of Escherichia coli acetohydroxyacid synthase isoenzyme II and characterization of three herbicide-insensitive forms. Biochem J 335:653–661, 1998. YT Lee, AK Chang, RG Duggleby. Eﬀect of mutagenesis at serine 653 of Arabidopsis thaliana acetohydroxyacid synthase on the sensitivity to imidazolinone and sulfonylurea herbicides. FEBS Lett 452:341–345, 1999. YT Lee, RG Duggleby. Mutagenesis studies on the sensitivity of Escherichia coli acetohydroxyacid synthase II to herbicides and valine. Biochem J 350:69– 73, 2000. G Jones, P Willett, RC Glen, AR Leach, R Taylor. Development and validation of a genetic algorithm for ﬂexible docking. J Molec Biol 267:727–748, 1997. JV Schloss, LM Ciskanik, DE Van Dyk. Origin of the herbicide binding site of acetolactate synthase. Nature 331:360–362, 1988. JV Schloss, A Aulabaugh. Acetolactate synthase and ketol-acid reductoisomerase: targets for herbicides obtained by screening and de novo design. Z Naturforsch 45c:544–551, 1990. S Mendel, T Elkayam, C Sella, V Vinogradov, M Vyazmensky, DM Chipman,

274

Duggleby et al.

Z Barak. Acetohydroxyacid synthase: a proposed structure for regulatory subunits supported by evidence from mutagenesis. J Molec Biol 307:465–477, 2001. 52. SS Pang, LW Guddat, RG Duggleby. Crystallization of the FAD-independent acetolactate synthase of Klebsiella pneumoniae. Acta Cryst D58:1237–1239, 2002.

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

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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.

REFERENCES 1.

2.

3.

4.

GD Hegeman. Synthesis of the enzymes of the mandelate pathway by Pseudomonas putida. I. Synthesis of enzymes by the wild type. J Bacteriol 91:1140– 1154, 1966. GD Hegeman. Synthesis of the enzymes of the mandelate pathway by Pseudomonas putida. II. Isolation and properties of blocked mutants. J Bacteriol 91: 1155–1160, 1966. GD Hegeman. Synthesis of the enzymes of the mandelate pathway by Pseudomonas putida. III. Isolation and properties of constitutive mutants. J Bacteriol 91:1161–1167, 1966. SL Rosenberg. Regulation of the mandelate pathway in Pseudomonas aeruginosa. J Bacteriol 108:1257–1269, 1971.

288 5.

6. 7. 8.

9. 10. 11.

12.

13.

14.

15.

16.

17.

18.

19.

Siegert et al. 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. DR Durham. Initial reactions involved in the dissimilation of mandelate by Rhodotorula graminis. J Bacteriol 160:778–780, 1984. GA Petsko, GL Kenyon, JA Gerlt, D Ringe, JW Kozarich. On the origin of enzymatic species. Trends Biochem Sci 18:372–376, 1993. S Bringer-Meyer, H Sahm. Pyruvate decarboxylase from Zymomonas mobilis: properties of the enzyme and catalysis of C–C bond formation. In: H Bisswanger, J Ullrich, eds. Biochemistry and Physiology of Thiamin Diphosphate Enzymes. Weinheim, Germany: VCH, 1991, pp. 123–132. M Pohl. Protein design on pyruvate decarboxylase (PDC) by site-directed mutagenesis. Adv Biochem Engin/Biotechnol 58:15–43, 1997. G Forlani, M Mantelli, E Nielsen. Biochemical evidence for multiple acetoinforming enzymes in cultured plant cells. Phytochemistry 50:255–262, 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. D Dobritzsch, S Konig, 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. F Dyda, W Furey, S Swaminathan, M Sax, B Farrenkopf, F Jordan. Catalytic centers in the thiamin diphosphate–dependent enzyme pyruvate decarboxylase at 2.4-A˚ resolution. Biochemistry 32:6165–6170, 1993. G Lu, D Dobritzsch, S Baumann, G Schneider, S Konig. The structural basis of substrate activation in yeast pyruvate decarboxylase. A crystallographic and kinetic study. Eur J Biochem 267:861–868, 2000. LS Polovnikova, MJ McLeish, EA Sergienko, JT Burgner, NL Anderson, F Kenyon, GL Kenyon, MS Hasson. Structural and kinetic analysis of catalysis by a thiamin diphosphate–dependent enzyme, benzoylformate decarboxylase. Biochemistry, 2002. (submitted for publication) G Schenk, FJ Leeper, R England, PF Nixon, RG Duggleby. The role of His113 and His114 in pyruvate decarboxylase from Zymomonas mobilis. Eur J Biochem 248:63–71, 1997. CY Huang, AK Chang, PF Nixon, RG Duggleby. Site-directed mutagenesis of the ionizable groups in the active site of Zymomonas mobilis pyruvate decarboxylase: eﬀect on activity and pH dependence. Eur J Biochem 268:3558– 3565, 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. AK Chang, PF Nixon, RG Duggleby. Aspartate-27 and glutamate-473 are

Substrate Specificity of BFD, PDC, and BAL

20.

21.

22.

23. 24. 25.

26. 27.

28.

29.

30.

31.

32.

33.

289

involved in catalysis by Zymomonas mobilis pyruvate decarboxylase. Biochem J 339:255–260, 1999. B Gonza´lez, R Vicun˜a. 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 Vicun˜a. 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.

290

Siegert et al.

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

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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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Scheme 9

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.

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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.

4.

5.

6.

7.

8.

9.

10.

11. 12. 13.

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

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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.

5. 6.

LJ Reed. Multienzyme complexes. Accounts Chem Res 7:40–46, 1974. LJ Reed, ML Hackert. Structure–function relationships in dihydrolipoamide acyltransferases. J Biol Chem 265:8971–8974, 1990. MS Patel, TE Roche. Molecular biology and biochemistry of pyruvate dehydrogenase complexes. FASEB J 4:3224–3233, 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. LJ Reed. A trail of research from lipoic acid to alpha-keto acid dehydrogenase complexes. J Biol Chem 276:38329–38336, 2001. RN Perham. Swinging arms and swinging domains in multifunctional enzymes: catalytic machines for multistep reactions. Annu Rev Biochem 69:961–1004, 2000.

328 7.

8.

9.

10.

11. 12.

13.

14.

15.

16.

17.

18.

19.

20.

Zhou et al. CY Maeng, MA Yazdi, XD Niu, HY Lee, LJ Reed. Expression, puriﬁcation, and characterization of the dihydrolipoamide dehydrogenase–binding protein of the pyruvate dehydrogenase complex from Saccharomyces cerevisiae. Biochemistry 33:13801–13807, 1994. S Gopalakrishnan, M Rahmatullah, GA Radke, S Powers-Greenwood, TE Roche. Role of protein X in the function of the mammalian pyruvate dehydrogenase complex. Biochem Biophys Res Commun 160:715–721, 1989. JC Neagle, JG Lindsay. Selective proteolysis of the protein X subunit of the bovine heart pyruvate dehydrogenase complex. Eﬀects on dihydrolipoamide dehydrogenase (E3) aﬃnity and enzymic properties of the complex. Biochem J 278:423–427, 1991. TE Roche, Y Hiromasa, A Turkan, X Gong, T Peng, X Yan, SA Kasten, H Bao, J Doug. Higher-domain-facilitated activated function and control of pyruvate dehydrogenase kinase and phosphatase isoform 1. Eur J Biochem. In Press 2002. RM Oliver, LJ Reed. Multienzyme complexes. In: ED Harris Jr, ed. Electron Microscopy of Proteins. London: Academic Press, 1992, pp. 1–48. T Wagenknecht, R Grassucci, D Schaak. Cryoelectron microscopy of frozenhydrated alpha-ketoacid dehydrogenase complexes from Escherichia coli. J Biol Chem 265:22402–22408, 1990. JK Stoops, TS Baker, JP Schroeter, SJ Kolodziej, XD Niu, LJ Reed. Threedimensional structure of the truncated core of the Saccharomyces cerevisiae pyruvate dehydrogenase complex determined from negative stain and cryoelectron microscopy images. J Biol Chem 267:24769–24775, 1992. T Wagenknecht, R Grassucci, GA Radke, TE Roche. Cryoelectron microscopy of mammalian pyruvate dehydrogenase complex. J Biol Chem 266:24650–24656, 1991. DJ DeRosier, RM Oliver, LJ Reed. Crystallization and preliminary structural analysis of dihydrolipoyl transsuccinylase, the core of the 2-oxoglutarate dehydrogenase complex. Proc Natl Acad Sci USA 68:1135–1137, 1971. CC Fuller, LJ Reed, RM Oliver, ML Hackert. Crystallization of a dihydrolipoyl transacetylase–dihydrolipoyl dehydrogenase subcomplex and its implications regarding the subunit structure of the pyruvate dehydrogenase complex from Escherichia coli. Biochem Biophys Res Commun 90:431–438, 1979. A Mattevi, G Obmolova, E Schulze, KH Kalk, AH Westphal, A de Kok, WG Hol. Atomic structure of the cubic core of the pyruvate dehydrogenase multienzyme complex. Science 255:1544–1550, 1992. A Mattevi, AJ Schierbeek, WG Hol. Reﬁned crystal structure of lipoamide dehydrogenase from Azotobacter vinelandii at 2.2-A˚ resolution. A comparison with the structure of glutathione reductase. J Mol Biol 220:975–994, 1991. A Mattevi, G Obmolova, JR Sokatch, C Betzel, WG Hol. The reﬁned crystal structure of Pseudomonas putida lipoamide dehydrogenase complexed with NAD+ at 2.45-A˚ resolution. Proteins 13:336–351, 1992. A Mattevi, G Obmolova, KH Kalk, WJ van Berkel, WG Hol. Three-dimensional structure of lipoamide dehydrogenase from Pseudomonas ﬂuorescens at 2.8-A˚

3D Structures of PDHCs

21.

22.

23.

24.

25.

26.

27.

28.

29. 30.

31.

32.

329

resolution. Analysis of redox and thermostability properties. J Mol Biol 230: 1200–1215, 1993. A Mattevi, G Obmolova, KH Kalk, AH Westphal, A de Kok, WG Hol. Reﬁned crystal structure of the catalytic domain of dihydrolipoyl transacetylase (E2p) from Azotobacter vinelandii at 2.6-A˚ resolution. J Mol Biol 230:1183–1199, 1993. 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. JDF Green, ED Laue, RN Perham, ST Ali, JR Guest. Three-dimensional structure of a lipoyl domain from the dihydrolipoyl acetyltransferase component of the pyruvate dehydrogenase multienzyme complex of Escherichia coli. J Mol Biol 248:328–343, 1995. SS Mande, S Sarfaty, MD Allen, RN Perham, WG Hol. Protein–protein interactions in the pyruvate dehydrogenase multienzyme complex: dihydrolipoamide dehydrogenase complexed with the binding domain of dihydrolipoamide acetyltransferase. Structure 4:277–286, 1996. YN Kalia, SM Brocklehurst, DS Hipps, E Appella, K Sakaguchi, RN Perham. The high-resolution structure of the peripheral subunit–binding domain of dihydrolipoamide acetyl-transferase from the pyruvate dehydrogenase multienzyme complex of Bacillus stearothermophilus. J Mol Biol 230:323–341, 1993. A Aevarsson, K Seger, S Turley, JR Sokatch, WG Hol. Crystal structure of 2-oxoisovalerate and dehydrogenase and the architecture of 2-oxo acid dehydrogenase multienzyme complexes. Nat Struct Biol 6:785–792, 1999. ZH Zhou, W Liao, RH Cheng, JE Lawson, DB McCarthy, LJ Reed, JK Stoops. Direct evidence for the size and conformational variability of the pyruvate dehydrogenase complex revealed by three-dimensional electron microscopy. The ‘‘breathing’’ core and its functional relationship to protein dynamics. J Biol Chem 276:21704–21713, 2001. T Izard, A Aevarsson, MD Allen, AH Westphal, RN Perham, A de Kok, WG Hol. Principles of quasi-equivalence and Euclidean geometry govern the assembly of cubic and dodecahedral cores of pyruvate dehydrogenase complexes. Proc Natl Acad Sci USA 96:1240–1245, 1999. W Chiu. Electron microscopy of frozen, hydrated biological specimens. Ann Rev Biophys Biophys Chem 15:237–257, 1986. J Dubochet, M Adrian, JJ Chang, JC Homo, J Lepault, AW McDowall, P Schultz. Cryoelectron microscopy of vitriﬁed specimens. Q Rev Biophys 21:128– 229, 1988. JK Stoops, RH Cheng, MA Yazdi, CY Maeng, JP Schroeter, U Klueppelberg, SJ Kolodziej, TS Baker, LJ Reed. On the unique structural organization of the Saccharomyces cerevisiae pyruvate dehydrogenase complex. J Biol Chem 272:5757–5764, 1997. ZH 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.

330

Zhou et al.

33. C-Y Maeng, MA Yazdi, LJ Reed. Stoichiometry of binding of mature and truncated forms of the dihydrolipoamide dehydrogenase-binding protein to the dihydrolipoamide acetyltransferase core of the pyruvate dehydrogenase complex from Saccharomyces cerevisiae. Biochemistry 35:5879–5882, 1996. 34. FH Pettit, LJ Reed. Pyruvate dehydrogenase complex from bovine kidney and heart. Methods Enzymol 89:376–386, 1982. 35. ZH Zhou, J He, J Jakana, JD Tatman, FJ Rixon, W Chiu. Assembly of VP26 in herpes simplex virus-1 inferred from structures of wild-type and recombinant capsids. Nat Struct Biol 2:1026–1030, 1995. 36. ZH Zhou, W Chiu, K Haskell, H Spears, Jr, J Jakana, FJ Rixon, LR Scott. Reﬁnement of herpesvirus B–capsid structure on parallel supercomputers. Biophys J 74:576–588, 1998. 37. Y Liang, EY Ke, ZH Zhou. IMIRS: high-resolution 3D reconstruction package integrated with a relational image database. J Struct Biol 137:292–304, 2002. 38. MA Robien, GM Clore, JG Omichinski, RN Perham, E Appella, K Sakaguchi, AM Gronenborn. Three-dimensional solution structure of the E3-binding domain of the dihydrolipoamide succinyltransferase core from the 2-oxoglutarate dehydrogenase multienzyme complex of Escherichia coli. Biochemistry 31: 3463–3471, 1992. 39. GA Carlson, K Hsiao, B Oesch, D Westaway, SB Prusiner. Genetics of prion infections. Trends Genet 7:61–65, 1991. 40. G Zaccai. How soft is a protein? A protein dynamics force constant measured by neutron scattering. Science 288:1604–1607, 2000. 41. E Wilson. Enzyme dynamics. Chem Engineer News 78:42–45, 2000. 42. CI Brandon, J Tooze. Introduction to Protein Structure. New York: Garland, 1991. 43. IA Balabin, JN Onuchic. Dynamically controlled protein tunneling paths in photosynthetic reaction centers. Science 290:114–117, 2000. 44. BW Lennon, CH Williams Jr, ML Ludwig. Twists in catalysis: alternating conformations of Escherichia coli thioredoxin reductase. Science 289:1190–1194, 2000. 45. Y Gu, ZH Zhou, DB McCarthy, LJ Reed, JK Stoops. 3D electron microscopy reveals the variable deposition and protein dynamics of the peripheral pyruvate dehydrogenase component about the core. Proc Natl Acad Sci USA 100:7015– 7020, 2003.

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

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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.

2.

3. 4.

5.

6.

7.

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

PDH Complex as a Molecular Machine

8.

9.

10.

11.

12.

13.

14. 15.

16.

17.

18.

19.

20.

341

interactions in the pyruvate dehydrogenase multienzyme complex: dihydrolipoamide dehydrogenase complexed with the binding domain of dihydrolipoamide acetyltransferase. Structure 4:277–286, 1996. IAD Lessard, RN Perham. Interaction of component enzymes with the peripheral subunit-binding domain of the pyruvate dehydrogenase multienzyme complex from Bacillus stearothermophilus: stoichiometry and speciﬁcity in selfassembly. Biochem J 306:727–733, 1995. IAD Lessard, C Fuller, RN Perham. Competitive interaction of component enzymes with the peripheral subunit-binding domain of the pyruvate dehydrogenase multienzyme complex of Bacillus stearothermophilus: kinetic analysis using surface plasmon resonance detection. Biochemistry 35:16863–16870, 1996. GJ Domingo, HJ Chauhan, IAD Lessard, C Fuller, RN Perham. Self-assembly and catalytic activity of the pyruvate dehydrogenase multienzyme complex from Bacillus stearothermophilus. Eur J Biochem 266:1136–1146, 1999. T Izard, A Ævarsson, MD Allen, AH Westphal, RN Perham, A de Kok, WGJ Hol. Principles of quasi-equivalence and Euclidean geometry govern the assembly of cubic and dodecahedral cores of pyruvate dehydrogenase complexes. Proc Natl Acad Sci USA 96:1240–1245, 1999. A Ævarsson, K Seger, S Turley, JR Sokatch, WGJ Hol. Crystal structure of 2oxoisovalerate dehydrogenase and the architecture of 2-oxo acid dehydrogenase multienzyme complexes. Nature Struct Biol 6:785–792, 1999. A Ævarsson, JL Chuang, RM Wynn, S Turley, DT Chuang, WGJ Hol. Crystal structure of human branched-chain a-ketoacid dehydrogenase and the molecular basis of multienzyme complex deﬁciency in maple syrup urine disease. Structure 8:277–291, 2000. LJ Reed. Multienzyme complexes. Acc Chem Res 7:40–46, 1974. NG Wallis, RN Perham. Structural dependence of post-translational modiﬁcation and reductive acetylation of the lipoyl domain of the pyruvate dehydrogenase multienzyme complex. J Mol Biol 236:209–216, 1994. LD Graham, LC Packman, RN Perham. Kinetics and speciﬁcity of reductive acylation of lipoyl domains from 2-oxo acid dehydrogenase multienzyme complexes. Biochemistry 28:1574–1581, 1989. DD Jones, HJ Horne, PA Reche, RN Perham. Structural determinants of posttranslational modiﬁcation and catalytic speciﬁcity for the lipoyl domains of the pyruvate dehydrogenase multienzyme complex of Escherichia coli. J Mol Biol 295:289–306, 2000. A Berg, AH Westphal, HJ Bosma, A de Kok. Kinetics and speciﬁcity of reductive acylation of wild-type and mutated lipoyl domains of 2-oxo-acid dehydrogenase complexes from Azotobacter vinelandii. Eur J Biochem 252:45–50, 1998. NG Wallis, MD Allen, RW Broadhurst, IAD Lessard, RN Perham. Recognition of a surface loop of the lipoyl domain underlies substrate channelling in the pyruvate dehydrogenase multienzyme complex. J Mol Biol 263:463–474, 1996. MJ Howard, HJ Chauhan, GJ Domingo, C Fuller, RN Perham. Protein–protein interaction revealed by NMR T2 relaxation experiments. The lipoyl domain and E1 component of the pyruvate dehydrogenase multienzyme complex of Bacillus stearothermophilus. J Mol Biol 295:1023–1047, 2000.

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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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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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mouse cDNA encoding a lipoic acid synthase that complements E. coli lipA mutant was isolated (50). Further studies will be required to clarify whether or not there is another lipoylation pathway in mammals corresponding to that found in E. coli. REFERENCES 1. 2.

3.

4.

5. 6.

7.

8.

9. 10.

11. 12.

13.

14.

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. K Fujiwara, K Okamura-Ikeda, Y Motokawa. Chicken liver H-protein, a component of the glycine cleavage system: amino acid sequence and identiﬁcation of the N-lipoyllysine residue. J Biol Chem 261:8836–8841, 1986. K Fujiwara, K Okamura-Ikeda, Y Motokawa. Expression of mature bovine Hprotein of the glycine cleavage system in Escherichia coli and in vitro lipoylation of the apoform. J Biol Chem 267:20011–20016, 1992. LJ Reed, FR Leach, M Koike. Studies on a lipoic acid–activating system. J Biol Chem 232:123–142, 1958. TW Morris, KE Reed, JE Cronan Jr. Identiﬁcation of the gene encoding lipoate-protein ligase A of Escherichia coli: molecular cloning and characterization of the lplA gene and gene product. J Biol Chem 269:16091–16100, 1994. DE Green, TW Morris, J Green, JE Cronan Jr, JR Guest. Puriﬁcation and properties of the lipoate protein ligase of Escherichia coli. Biochem J 309:853– 862, 1995. K Fujiwara, K Okamura-Ikeda, Y Motokawa. Puriﬁcation and characterization of lipoyl-AMP:N q-lysine lipoyltransferase from bovine liver mitochondria. J Biol Chem 269:16605–16609, 1994. K Fujiwara, K Okamura-Ikeda, Y Motokawa. Cloning and expression of a cDNA encoding bovine lipoyltransferase. J Biol Chem 272:31974–31978, 1997. K Fujiwara, S Takeuchi, K Okamura-Ikeda, Y Motokawa. Puriﬁcation, characterization, and cDNA cloning of lipoate-activating enzyme from bovine liver. J Biol Chem 276:28819–28823, 2001. JN Tsunoda, KT Yasunobu. Mammalian lipoic acid activating enzyme. Arch Biochem Biophys 118:395–401, 1967. CM Smith, J Bryla, JR Williamson. Regulation of mitochondrial a-ketoglutarate metabolism by product inhibition at a-ketoglutarate dehydrogenase. J Biol Chem 249:1497–1505, 1974. DA Vessey, E Lau, M Kelley. Isolation and sequencing of cDNAs for XL-I and XL-III forms of bovine liver xenobiotic-metabolizing medium-chain fatty acid: CoA ligase. J Biochem Mol Toxicol 14:11–19, 2000. DA Vessey, M Kelly, E Lau, SZ Zhang. Monovalent cation eﬀects on the

Protein Lipoylation in Mammals

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

359

activity of the xenobiotic/medium-chain fatty acid:CoA ligases are substrate speciﬁc. J Biochem Mol Toxicol 14:162–168, 2000. K Fujiwara, K Okamura-Ikeda, Y Motokawa. Lipoylation of acyltransferase components of a-ketoacid dehydrogenase complexes. J Biol Chem 271:12932– 12936, 1996. TA Griﬃn, KS Lau, DT Chuang. Characterization and conservation of the inner E2 core domain structure of branched-chain a-keto acid dehydrogenase complex from bovine liver: construction of a cDNA encoding the entire transacylase (E2b) precursor. J Biol Chem 263:14008–14014, 1988. S Matuda, K Nakano, S Ohta, M Shimura, T Yamanaka, S Nakagawa, K Titani, T Miyata. Molecular cloning of dihydrolipoamide acetyltransferase of the rat pyruvate dehydrogenase complex: sequence comparison and evolutionary relationship to other dihydrolipoamide acyltransferases. Biochim Biophys Acta 1131:114–118, 1992. TJ Thekkumkara, L Ho, ID Wexler, G Pons, T-C Liu, MS Patel. Nucleotide sequence of a cDNA for the dihydrolipoamide acetyltransferase component of human pyruvate dehydrogenase complex. FEBS Lett 240:45–48, 1988. X-D Niu, KS Browning, RH Behal, LJ Reed. Cloning and nucleotide sequence of the gene for dihydrolipoamide acetyltransferase from Saccharomyces cerevisiae. Proc Natl Acad Sci USA 85:7546–7550, 1988. PE Stephens, MG Darlison, HM Lewis, JR Guest. The pyruvate dehydrogenase complex of Escherichia coli K12: nucleotide sequence encoding the dihydrolipoamide acetyltransferase component. Eur J Biochem 133:481–489, 1983. R Hanemaaijer, A Janssen, A de Kok, C Veeger. The dihydrolipoyltransacetylase component of the pyruvate dehydrogenase complex from Azotobacter vinelandii: molecular cloning and sequence analysis. Eur J Biochem 174:593– 599, 1988. K Nakano, S Matuda, T Yamanaka, H Tsubouchi, S Nakagawa, K Titani, S Ohta, T Miyata. Puriﬁcation and molecular cloning of succinyltransferase of the rat a-ketoglutarate dehydrogenase complex: absence of a sequence motif of the putative E3- and/or E1-binding site. J Biol Chem 266:19013–19017, 1991. ME Spencer, MG Darlison, PE Stephens, IK Duckenﬁeld, JR Guest. Nucleotide sequence of the sucB gene encoding the dihydrolipoamide succinyltransferase of Escherichia coli K12 and homology with the corresponding acetyltransferase. Eur J Biochem 141:361–374, 1984. AH Westphal, A de Kok. The 2-oxoglutarate dehydrogenase complex from Azotobacter vinelandii: 2. Molecular cloning and sequence analysis of the gene encoding the succinyltransferase component. Eur J Biochem 187:235–239, 1990. K Fujiwara, K Okamura-Ikeda, Y Motokawa. cDNA sequence, in vitro synthesis, and intramitochondrial lipoylation of H-protein of the glycine cleavage system. J Biol Chem 265:17463–17467, 1990. K Fujiwara, K Okamura-Ikeda, K Hayasaka, Y Motokawa. The primary structure of human H-protein of the glycine cleavage system deduced by cDNA cloning. Biochem Biophys Res Commun 176:711–716, 1991. H Koyata, K Hiraga. The glycine cleavage system: structure of a cDNA en-

360

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

Fujiwara et al. coding human H-protein, and partial characterization of its gene in patients with hyperglycinemias. Am J Hum Genet 48:351–361, 1991. M Yamamoto, H Koyata, C Matsui, K Hiraga. The glycine cleavage system: occurrence of two types of chicken H-protein mRNAs presumably formed by the alternative use of the polyadenylation consensus sequences in a single exon. J Biol Chem 266:3317–3322, 1991. Y Kim, DJ Oliver. Molecular cloning, transcriptional characterization, and sequencing of cDNA encoding the H-protein of the mitochondrial glycine decarboxylase complex in peas. J Biol Chem 265:848–853, 1990. D Macherel, M Lebrum, J Gagnon, M Neuburger, R Douce. cDNA cloning, primary structure and gene expression for H-protein, a component of the glycine-cleavage system (glycine decarboxylase) of pea (Pisum sativum) leaf mitochondria. Biochem J 268:783–789, 1990. LT Stauﬀer, PS Steiert, JG Steiert, GV Stauﬀer. An Escherichia coli protein with homology to the H-protein of the glycine cleavage enzyme complex from pea and chicken liver. DNA Seq 2:13–17, 1991. K Okamura-Ikeda, Y Ohmura, K Fujiwara, Y Motokawa. Cloning and nucleotide sequence of the gcv operon encoding the Escherichia coli glycinecleavage system. Eur J Biochem 216:539–548, 1993. KS Lau, TA Griﬃn, C-WC Hu, DT Chuang. Conservation of primary structure in the lipoyl-bearing and dihydrolipoyl dehydrogenase–binding domains of mammalian branched-chain a-keto acid dehydrogenase complex: molecular cloning of human and bovine transacylase (E2) cDNAs. Biochemistry 27:1972–1981, 1988. K Ono, M Hakozaki, T Suzuki, T Mori, H Hata, H Kochi. cDNA cloning of the chicken branched-chain a-keto acid dehydrogenase complex: chickenspeciﬁc residues of the acyltransferase aﬀect the overall activity and the interaction with the dehydrogenase. Eur J Biochem 268:727–736, 2001. 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. JDF Green, ED Laue, RN Perham, ST Ali, JR Guest. Three-dimensional structure of a lipoyl domain from the dihydrolipoyl acetyltransferase component of the pyruvate dehydrogenase multienzyme complex of Escherichia coli. J Mol Biol 248:328–343, 1995. A Berg, J Vervoort, A de Kok. Three-dimensional structure in solution of the N-terminal lipoyl domain of the pyruvate dehydrogenase complex from Azotobacter vinelandii. Eur J Biochem 244:352–360, 1997. A Berg, J Vervoort, A de Kok. Solution structure of the lipoyl domain of the 2-oxoglutarate dehydrogenase complex from Azotobacter vinelandii. J Mol Biol 261:432–442, 1996. PM Ricaud, MJ Howard, EL Roberts, RW Broadhurst, RN Perham. Threedimensional structure of the lipoyl domain from the dihydrolipoyl succinyltransferase component of the 2-oxoglutarate dehydrogenase multienzyme complex of Escherichia coli. J Mol Biol 264:179–190, 1996.

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40. S Pares, C Cohen-Addad, L Sieker, M Neuburger, R Douce. X-ray structure determination at 2.6-A˚ resolution of a lipoate-containing protein: The H-protein of the glycine decarboxylase complex from pea leaves. Proc Natl Acad Sci USA 91:4850–4853, 1994. 41. NG Wallis, RN Perham. Structural dependence of post-translational modiﬁcation and reductive acetylation of the lipoyl domain of the pyruvate dehydrogenase multienzyme complex. J Mol Biol 236:209–216, 1994. 42. K Fujiwara, M Suzuki, Y Okumachi, K Okamura-Ikeda, T Fujiwara, E Takahashi, Y Motokawa. Molecular cloning, structural characterization and chromosomal localization of human lipoyltransferase gene. Eur J Biochem 260: 761–767, 1999. 43. K Okamura-Ikeda, K Fujiwara, M Yamamoto, K Hiraga, Y Motokawa. Isolation and sequence determination of cDNA encoding T-protein of the glycine cleavage system. J Biol Chem 266:4917–4921, 1991. 44. J Peinado, H Sies, TPM Akerboom. Hepatic lipoate uptake. Arch Biochem Biophys 273:389–395, 1989. 45. SM Grassl. Human placental brush-border membrane Na+-biotin cotransport. J Biol Chem 267:17760–17765, 1992. 46. PD Prasad, H Wang, R Kekuda, T Fujita, Y-J Fei, LD Devoe, FH Leibach, V Ganapathy. Cloning and functional expression of a cDNA encoding a mammalian sodium-dependent vitamin transporter mediating the uptake of pantothenate, biotin, and lipoate. J Biol Chem 273:7501–7506, 1998. 47. KE Reed, JE Cronan Jr. Lipoic acid metabolism in Escherichia coli: sequencing and functional characterization of the lipA and lipB genes. J Bacteriol 175: 1325–1336, 1993. 48. JR Miller, RW Busby, SW Jordan, J Cheek, TF Henshaw, GW Ashley, JB Broderick, JE Cronan Jr., MA Marletta. Escherichia coli LipA is a lipoyl synthase: in vitro biosynthesis of lipoylated pyruvate dehydrogenase complex from octanoyl-acyl carrier protein. Biochemistry 39:15166–15178, 2000. 49. TW Morris, KE Reed, JE Cronan Jr. Lipoic acid metabolism in Escherichia coli: the lplA and lipB genes deﬁne redundant pathways for ligation of lipoyl groups to apoprotein. J Bacteriol 177:1–10, 1995. 50. T Morikawa, R Yasuno, H Wada. Do mammalian cells synthesize lipoic acid? Identiﬁcation of a mouse cDNA encoding a lipoic acid synthase located in mitochondria. FEBS Lett 498:16–21, 2001.

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 (<3 nM). PDK2 and PDK3 achieve rapid initial rates when only a few E1 are bound to the E2 60mer and, as indicated earlier, these kinases also sustain high rates with dilute complexes (Y. Hiromasa and T.E. Roche, unpublished). We propose that the capacity, documented earlier, for bifunctional binding of these kinases by the L2 domain underpins the E2 60mer providing rapid access to successive E2-bound E1. Speciﬁcally, continuous access to E1 is suggested to result from ‘‘hand-over-hand’’ transfer on the surface of E2 60mer via a kinase dimer bound by two lipoyl domains, randomly letting go of one lipoyl domain and then rebinding a second mobile lipoyl domain faster than letting go from the singly held state (54). Each of the lipoyl domains of E2 is concentrated within the exterior of the complex at >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 (<10%) in the assembled complex undergoes acetylation (70,71). Near-maximal stimulation of the kinase activity is attained with very low product-to-substrate ratios—0.1 NADH/NAD+ and 0.2 acetyl-CoA/CoA (74). The greater extended reach of the reduced and acetylated lipoyl group over that of the oxidized prosthetic group (Fig. 4) likely bestows the ability for interactions to induce activating conformations within the kinase active site. The primary kinases in kidney medulla are PDK2 and PDK3. PDK2 is the most responsive human kinase to product stimulation (22,56). The much stronger binding of PDK2 upon reduction of the lipoyl group of dimeric GST-L2 but not L2 (see earlier) is consistent with additional interactions associated with bifunctional binding that elicit a conformational change in the PDK structure that elevates kinase catalysis. Under conditions of physiological salts (particularly high K+ ion), the full set of regulatory properties of PDK2 are observed in the presence of E2 (56; X. Yan, H. Bao, S.A. Kasten, and T.E. Roche, unpublished work). Elevation of ADP and phosphate signals a low-energy state; ample pyruvate indicates the abundance of glucose-linked substrates for meeting this energy demand. These energy-demanding conditions forcefully inhibit kinase inactivation of PDC activity. The marked elevation of PDK2 activity by E2 transforms PDK2 from being insensitive to pyruvate or dichloroacetate (DCA) to being potently inhibited by these allosteric eﬀectors (55). Indeed, synergistic inhibition by pyruvate (or DCA) and ADP is produced via these inhibitors’ binding to PDK2.ADP (and PDK2.ATP) but not to the free PDK2. Phosphate inhibition of PDK2 is also favored and acts together with ADP and pyruvate in reducing PDK2 activity. Based on kinetics and the common requirements for E2 and elevated K+ ion, eﬀector control of PDK2 activity appears to be mechanistically consolidated by inhibitors slowing down and activators speeding up a rate-limiting ADP dissociation (1,55,56; X. Yan, H. Bao, S.A. Kasten, and T.E. Roche, unpublished work). In contrast to PDK2, PDK3 is only weakly inhibited by pyruvate. However, the combination of phosphate and ADP inhibits PDK3 activity in more than an additive manner, indicating that PDK3 activity is impaired when the level of ATP decreases (56). Human PDK3 activity is increased by product stimulation only when countering the eﬀect of an inhibitor (e.g., by ADP) or in counteracting other conditions that lower PDK3 activity (56). As an example of the latter, PDK3 activity is increased several-fold when only the poorly activating L1 domain of E2 has a reactive lipoyl group available for undergoing reductive acetylation. The gain in the capacity for PDK3 activity to be enhanced by reductive acetylation in the presence of ADP and phosphate appears to be the primary basis for the higher

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fractional stimulation of bovine kidney kinases observed in the presence of these inhibitors (69). D. Binding Specificity and Regulation of PDK4 In attempting to express human PDK4 (J. Dong, L. Hu, and T.E. Roche, unpublished work), we found that the hydrophobic C-terminal Val-Ala-Met sequence triggered degradation of PDK4 recombinantly expressed in E. coli. To prevent this, we ﬁrst added a Gly-Glu-Glu amino acid sequence. Subsequently, we produced unmodiﬁed PDK4 by using an E. coli strain that lacked the C1pP proteosome. In comparing properties of the GEE-modiﬁed and unmodiﬁed PDK4, we found that only the unmodiﬁed PDK4 was stimulated by NADH and acetyl-CoA. This suggests that the C-terminal region of PDKs is important for the lipoyl-mediated product stimulation. Interestingly, PDK4 is preferentially bound by the L1 domain of E2 and the L3 domain of E3BP (Fig. 1), and the reduction and acetylation of the lipoyl groups of the L1 and L3 domains enhance PDK4 activity. PDK1 binds to both the L1 and L2 domains of E2 (Fig. 1) (J.C. Baker, X. Yan, and T.E. Roche, unpublished). PDK1 is by far the most eﬀective in phosphorylating the third phosphorylation site on E1 (75,76); however, contrary to the conclusion of these studies, we have found limited phosphorylation of this site by the other PDKs (S.A. Kasten and T.E. Roche, unpublished). IV. E2-MEDIATED CA2+ ACTIVATION OF PDP1 Elevation of intramitochondrial free Mg2+ due to diminished binding by ATP serves as a secondary signal of a cellular energy deﬁcit. Exercise, growth, and many other neural and hormone-induced cellular transitions that demand energy elicit a rise in cellular Ca2+ in the cytoplasm that eventuates in an increase in intramitochondrial Ca2+ (9). PDP1 responds to both these crucial indicators of energy need and acts to increase the proportion of active PDC. PDP1 activity is directly dependent on and highly sensitive to the physiological range of Mg2+, which is not thought to vary outside of 0.4–1.5 mM in normally functioning mitochondria. PDP1 activity is also ampliﬁed by physiological increases (from submicromolar to micromolar) in free Ca2+. PDP1 is a heterodimer (24,77) consisting of a 52-kDa catalytic subunit (23), PDP1c, and a 96-kDa regulatory subunit, PDP1r (25). In the absence of PDP1r, PDP1c has a low Km of 0.4 mM for Mg2+ (23). However, in holo-PDP1, PDP1r promulgates an increase in the Km for Mg2+ to 2 mM in the presence of saturating Ca2+ and to 3.5 mM in the absence of Ca2+ (27). Binding of spermine reverses the eﬀect of the PDP1r subunit by returning the Km of holo-PDP1 for Mg2+ to the lower level (0.5 mM). The Ca2+- and

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spermine-induced changes in responsiveness to Mg2+ evoke marked changes in PDP1 activity at physiological levels of intramitochondrial free Mg2+. E2 greatly enhances PDP1 activity through Ca2+-dependent binding of PDP1 or just PDP1c to the L2 domain of E2 (58,78). With saturating Mg2+, E2 plus Ca2+ increases the rates of dephosphorylation by PDP1 by 10-fold and by PDP1c by 6-fold. Larger increases are promoted by E2/Ca2+ when phosphatase activity is compared with free and E2-bound phosphorylated-E1 with subsaturating levels of Mg2+ or<1 lM E1. The activation of PDP1 or PDP1c by E2 involves highly speciﬁc Ca2+-dependent binding to the L2 domain of E2 by an interaction that requires both precise surface structure of the L2 domain and the lipoyl prosthetic group (58,78). We have employed a set of over 30 mutant L2 domains and substitutions within the lipoyl-lysine prosthetic group to evaluate the speciﬁc structure by which L2 binds PDP1 and PDP1c (78). We have found two major regions. The ﬁrst encompasses the lipoyl prosthetic group and neighboring residues (Fig. 3A, p or pp residues; see legend). Binding is markedly reduced by substitutions for Ala172 and Asp173 as well as Leu140. Binding was lost with nonlipoylated L2 or following substitution by any of a series of amino acid for lipoylated Lys173. However, binding was fully retained upon replacing the lipoyl group with an octanoyl group. The latter ﬁnding suggests that the lipoyl cofactor interacts within an extended hydrophobic pocket in the surface of PDP1c. In contrast to just an octanoyl group is being eﬀective, the dithiolane ring of the lipoyl prosthetic group makes a major contribution to L2 binding to E1 (79) and, as indicated earlier, the 8-thiol is pivotal for L2 activating PDK3. The L1 domain of E2 has equivalent structure at its lipoylated end, so other aspects of L2’s structure are required to explain the high speciﬁcity for PDP1 binding to E2’s inner lipoyl domain. At the other end of the L2 domain, mutation of glutamates 162, 179, and 182 and glutamine 181 markedly hinders binding of PDP1 and PDP1c (78). Substitution of alanine or glutamine for Glu182 prevented binding of PDP1 and PDP1c to the modiﬁed L2 domains. A distinct pocket, Fig. 3A, is formed by this cluster of residues. Substitution of any of the acidic residues increases the stability of the domain, probably due to reduction of the mutual repulsion of the similarly charged acidic residues. Full or partial removal of the this electrostatic repulsion may contribute to the Ca2+-dependent binding of PDP1c by this region. An attractive hypothesis is that Ca2+ forms a bridge by interacting in this region. Mutation of Glu179 did not change the concentration dependence for Ca2+; this could not be evaluated for Glu182 because substitutions (Ala or Glu) at this residue completely eliminated binding. The conversion of the Val-Gln residues located between Glu179 and Glu182 in L2 to the Ser-Leu sequence connecting equivalent acidic residues in L1 eﬀectively hindered binding of PDP1 to L2 (76). This dual substitution had

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a lesser but signiﬁcant eﬀect on binding of PDP1c. In contrast, this linked mutation did not reduce the use of L2 in the E1 reaction, even though Glu179 is a key speciﬁcity residue for E1 (79). Five other single-site substitutions (A174S and R196Q at the lipoylated end of L2, D213N and Y220A in the Cterminal lobe, and the Y129A in the N-terminal segment) caused a greater reduction in L2’s binding to PDP1 than to PDP1c, suggesting that the PDP1r subunit of PDP1 directly or indirectly contributes to a more precise interaction with L2. Normally when Ca2+ is bound with a near -1-lM aﬃnity, Ca2+ binding involves an EF-hand structure, but based on the structure of L2 and the sequence of PDP1c, no EF-hand is apparent. Isothermal titration calorimetry and direct Ca2+-binding measurements revealed that Ca2+ does not bind to either L2 or PDP1c, alone (A. Turkan and T.E. Roche, unpublished work). This indicates that Ca2+ either directly engages in forming a bridge between PDP1c and L2 or enhances binding by a capture mechanism. Because the association between L2 and PDP1c involves both a domain-aided hydrophobic interaction by the extended lipoyl-lysine cofactor at one end of L2 and electrostatic interactions by a cluster of residues at the opposite end of the domain (78), it seems likely that these essential regions act in concert. An attractive prospect is that these interacting regions of L2 work together to promote a substantial conformation transition in PDP1c that fosters and stabilizes a tight Ca2+ binding. Detailed structures, probably dependent on crystallizing these complexes, will be required to elucidate how this unusual and highly activating complex is formed. V. SUMMARY We have emphasized evidence that the E2-E3BP structure in mammalian PDC is fabricated by the integration of the C-terminal domain of E3BP into the dodecahedron structure formed by the structurally related C-terminal domain of E2. With a focus on the kinase isoforms PDK2 and PDK3 and on PDP1 and its catalytic subunit, PDP1c, we have considered results that manifest how interactions with the ﬂexibly held outer domains of the E2 assemblage have a predominant role in determining the relative rates of phosphorylation and dephosphorylation of the E1 component. These interactions, in part, involve speciﬁc binding to the L2 domain of E2 and act to up-regulate the activities of these enzymes and mediate or alter their responsiveness to regulatory eﬀectors. A common feature contributing to greatly enhanced activity of the regulatory enzymes is that their ﬂexible binding at the E2 surface facilitates greatly enhanced access to their E1 substrates. The highly favored interactions of PDK3 and PDP1 with the L2 domain of E2 were found to require extensive and well-separated regions of the surface of this lipoyl

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domain acting in concert with the lipoyl prosthetic group. E2 mediates stimulation of kinase activities, particularly PDK2, by reduction and acetylation of the lipoyl cofactor when consumption of fatty acids leads to elevation of NADH and acetyl-CoA. Via the L2 domain, E2 directly participates in the Ca2+-dependent activation of PDP1 and its catalytic subunit, PDP1c. E2 also alters kinase and phosphatase activities by altering the response to ligands that bind directly to the regulatory enzymes (e.g., enhance pyruvate inhibition of PDK2, alter the Km of PDP1 for Mg2+). Therefore, the E2 oligomer not only plays a pivotal integrating role in binding the catalytic components and consolidating the catalytic reactions of the complex, but also facilitates and helps regulate the kinase/phosphatase control of PDC activity.

ACKNOWLEDGMENTS This work was supported by the National Institutes of Health Grant DK18320 and by the Kansas Agriculture Experiment Station—contribution 03—J. ABBREVIATIONS PDC, pyruvate dehydrogenase complex E1, pyruvate dehydrogenase component E2, dihydrolipoyl acetyltransferase component L1 domain, NH2-lipoyl domain of E2 L2 domain, interior lipoyl domain of E2 PDK, pyruvate dehydrogenase kinase PDK1, PDK2, PDK3 and PDK4, PDK isoforms 1, 2, 3, and 4 PDP, pyruvate dehydrogenase phosphatase PDP1 and PDP2, PDP isoforms 1 and 2 PDP1c, catalytic subunit of PDP1 PDP1r, regulatory subunit of PDP1 E3, dihydrolipoyl dehydrogenase; E3BP, E3-binding protein L3, N-terminal lipoyl domain of E3BP GST, glutathione-S-transferase DCA, dichloroacetate AUC, analytical ultracentrifugation

REFERENCES 1.

TE Roche, J Liu, S Ravindran, J Baker, D Wang. Role of the E2 core in the dominant mechanisms of regulatory control of mammalian pyruvate dehydro-

E2-Facilitated Kinase/Phosphatase Function

2.

3.

4.

5.

6.

7. 8. 9.

10.

11.

12.

13. 14.

15.

16.

381

genase complex. In: MS Patel, TE Roche, RA Harris, eds. Alpha-Keto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996, pp 33–52. TE Roche, DJ Cox. Multifunctional 2-Oxo acid dehydrogenase complexes. L. Agius, HSA Sherratt eds. Channeling in Intermediary Metabolism. London: Portland Press, 1996, pp 156–132. TE Roche, JC Baker, X Yan, Y Hiromasa, X Gong, T Peng, J Dong, A Turkan, SA Kasten. Distinct regulatory properties of pyruvate dehydrogenase kinase and phosphatase isoforms. Prog Nucleic Acid Res Mol Biol 70:33–75, 2001. TE Roche, Y Hiromasa, A Turkan, X Gong, T Peng, X Yan, SA Kasten, H Bao, J Dong. Lipoyl domain–facilitated activated function and control of pyruvate dehydrogenase kinases and phosphatase isoform 1. Eur J Biochem 270:1050– 1056, 2003. PJ Randle, DA Priestman. Shorter-term and longer-term regulation of pyruvate dehydrogenase kinases. In: MS Patel, TE Roche, RA Harris eds. AlphaKeto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996, pp 151– 161. MC Sugden, MJ Holness. Hormonal and nutritional maturation of PDHC activity status. In: MS Patel, TE Roche, RA Harris eds. Alpha-Keto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996, pp 163–176. RA Harris, B Huang, P Wu. Control of pyruvate dehydrogenase kinase gene expression. Advan Enzyme Regul 41:269–288, 2001. MC Sugden, K Bulmer, MJ Holness. Fuel-sensing mechanisms integrating lipid and carbohydrate utilization. Biochem Soc Trans 29:272–278, 2001. RM Denton, JG McCormack, GA Rutter, P Burnett, NJ Edgell, SK Moule, TA Diggle. The hormonal regulation of pyruvate dehydrogenase complex. Advan Enzyme Regul 36:183–198, 1996. R Gudi, MM Bowker-Kinley, NY Kedishvili, Y Zhao, KM Popov. Diversity of the pyruvate dehydrogenase kinase gene family in humans. J Biol Chem 270:28989–28994, 1995. J Rowles, SW Scherer, T Xi, M Majer, DC Nickle, JM Rommens, KM Popov, RA Harris, NL Riebow, J Xia, L-C Tsui, C Bogardus, M Prochazka. Cloning and characterization of PDK4 on 7q21.3 encoding a fourth pyruvate dehydrogenase kinase isozyme in human. J Biol Chem 271:22376–22382, 1966. B Huang, R Gudi, P Wu, RA Harris, J Hamilton, KM Popov. Isozymes of pyruvate dehydrogenase phosphatase. DNA-derived amino acid sequences, expression, and regulation. J Biol Chem 273:17680–17688, 1998. M Bower-Kinley, KM Popov. Evidence that pyruvate dehydrogenase kinase belongs to the ATPase/kinase superfamily. Biochem J 344:47–53, 1999. RM Wynn, JL Chaung, CD Cote, DT Chuang. Tetrameric assembly and conservation in the ATP-binding domain of rat branched-chain alpha-ketoacid dehydrogenase kinase. J Biol Chem 275:30512–30519, 2000. CN Steussy, KM Popov, MM Bowker-Kinley, RB Sloan, RA Harris, JA Hamilton. Structure of pyruvate dehydrogenase kinase. Novel folding pattern for a serine protein kinase. J Biol Chem 276:37443–37450, 2001. M Machius, JL Chaung, RM Wynn, DR Tomchick, DT Chuang. Structure of

382

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27. 28. 29.

30.

31.

Roche et al. rat BCKD kinase: nucleotide-induced domain communication in a mitochondrial protein kinase. Proc Nat Acad Sci USA 98:11218–11223, 2001. ML Pratt, TE Roche. Mechanism of pyruvate inhibition of kidney pyruvate dehydrogenase kinase and synergistic inhibition by pyruvate and ADP. J Biol Chem 254:7191–7196, 1979. P Wu, J Sato, Y Zhao, Ny Kedishvili, Y Shimomura, DW Crabb. Starvation and diabetes increase the amount of pyruvate dehydrogenase kinase isoform 4 in rat heart. Biochem J 329:197–201, 1998. P Wu, PV Blair, J Sato, J Jaskiewicz, KM Popov, RA Harris. Starvation increases the amount of pyruvate dehydrogenase kinase in several mammalian tissues. Arch Biochem Biophys 381:1–7, 2000. MJ Holness, A Kraus, RA Harris, MC Sugden. Targeted upregulation of pyruvate dehydrogenase kinase (PDK)-4 in slow-twitch skeletal muscle underlies stable modiﬁcation of the regulatory characteristics of PDK induced by fat feeding. Diabetes 49:775–781, 2000. P Wu, JM Peters, RA Harris. Adaptive increase in pyruvate dehydrogenase kinase 4 during starvation is mediated by peroxisome proliferator-activated receptor a. Biochem Biophys Res Commun 287:391–396, 2001. MM Bowker-Kinley, WI Davis, P Wu, RA Harris, KM Popov. Evidence for existence of tissue-speciﬁc regulation of the mammalian pyruvate dehydrogenase complex. Biochem J 329:191–196, 1998. JE Lawson, X-D Niu, KS Browning, HL Trong, J Yan, LJ Reed. Molecular cloning and expression of the catalytic subunit of bovine pyruvate dehydrogenase phosphatase and sequence similarity with protein phosphatase 2C. Biochemistry 32:8987–8993, 1993. WM Teague, FH Pettit, T-L Wu, SR Silberman, LJ Reed. Puriﬁcation and properties of pyruvate dehydrogenase phosphatase from bovine heart and kidney. Biochemistry 21:5585–5592, 1982. JE Lawson, SH Park, AR Mattison, J Yan, LJ Reed. Cloning, expression and properties of the regulatory subunit of bovine pyruvate dehydrogenase phosphatase. J Biol Chem 272:31625–31629, 1997. Z Damuni, JS Humphreys, LJ Reed. Stimulation of pyruvate dehydrogenase phosphatase activity by polyamines. Biochem Biophys Res Commun 124:95– 99. J Yan, JE Lawson, LJ Reed. Role of the regulatory subunit of bovine pyruvate dehydrogenase phosphatase. Proc Natl Acad Sci USA 93:4953–4956, 1996. LJ Reed, Z Damuni. Mitochondrial protein phosphatases. Adv Prot Phosphatases 4:59–76, 1987. LJ Reed, JE Lawson, X-D Niu, J Yan. Pyruvate dehydrogenase phosphatase. In: MS Patel, TE Roche, RA Harris eds. Alpha-Keto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996, pp 131–138. AP Thomas, RM Denton. Use of toluene permeabilized mitochondria to study the regulation of adipose tissue pyruvate dehydrogenase in situ. Biochem J 238:93–101, 1986. J Larner, LC Huang, S Suzuki, G Tang, C Zhang, CFW Schwartz, G Romero,

E2-Facilitated Kinase/Phosphatase Function

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

383

L Luttrell, AS Kennington. Insulin mediators and control of the pyruvate dehydrogenase complex. Ann NY Acad Sci 573:297–305, 1989. M Caruso, MA Maitan, G Bifulco, C Miele, G Vigliotta, F Oriente, P Formisano, F Beguinot. Activation and mitochondrial translocation of protein kinase C delta are necessary for insulin stimulation of pyruvate dehydrogenase complex activity in muscle and liver cells. J Biol Chem 276:45088–45097, 2001. TJ Thekkumkara, L Ho, ID Wexler, G Pons, T-C Lui, MS Patel. Nucleotide sequence of cDNA of the dihydrolipoyl acetyltransferase component of human pyruvate dehydrogenase complex. FEBS Lett 240:45–48, 1988. RA Harris, MM Bowker-Kinley, P Wu, J Jeng, KM Popov. Dihydrolipoamide dehydrogenase–binding protein of the human pyruvate dehydrogenase complex. DNA-derived amino acid sequence, expression, and reconstitution of the pyruvate dehydrogenase complex. J Biol Chem 272:19746–19751, 1997. M Rahmatullah, S Gopalakrishnan, PC Andrews, CL Chang, GA Radke, TE Roche. Subunit associations in the mammalian pyruvate dehydrogenase complex: structure and role of protein X and the pyruvate dehydrogenase component–binding domain of the dihydrolipoyl transacetylase component. J Biol Chem 264:2221–2227, 1989. M Rahmatullah, GA Radke, PC Andrews, TE Roche. Changes in the core of the mammalian-pyruvate dehydrogenase complex upon selective removal of the lipoyl domain from the transacetylase component but not from the protein X component. J Biol Chem 265:14512–14517, 1990. A Mattevi, G Obmolova, E Schuze, KH Kalk, A Westphal, A de Kok, WGJ Hol. The atomic structure of the cubic core of the pyruvate dehydrogenase complex. Science 255:1544–1550, 1992. A Mattevi, G Obmolova, KH Lalk, A Teplyakov, WGJ Hol. Crystallographic analysis of substrate binding and catalysis in dihydrolipoyl tansacetylase (E2p). Biochemistry 32:3887–3901, 1993. M Rahmatullah, S Gopalakrishnan, GA Radke, TE Roche. Domain structure of the dihydrolipoyl transacetylase and the protein X components of mammalian pyruvate dehydrogenase complex—selective cleavage by protease. Arg C J Biol Chem 264:1245–1251, 1989. SL Powers-Greenwood, M Rahmatullah, GA Radke, TE Roche. Separation of protein X from the dihydrolipoyl transacetylase component of the mammalian pyruvate dehydrogenase complex and role of protein X. J Biol Chem 264:3655– 3657, 1989. S Gopalakrishnan, M Rahmatullah, GA Radke, SL Powers-Greenwood, TE Roche. Role of protein X in the function of mammalian pyruvate dehydrogenase complex. Biochem Biophys Res Commun 160:715–721, 1989. JE Lawson, RH Behal, LJ Reed. Disruption and mutation of the Saccharomyces cervvisiae PDX1 gene encoding the protein X component of the pyruvate dehydrogenase complex. Biochemistry 30:2834–2839, 1991. SJ Sanderson, SS Khan, RG McCartney, CM Miller, GJ Lindsay. Reconstitution of mammalian pyruvate dehydrogenase and 2-oxoglutarate dehydrogenase complex: analysis of protein X involvement and interaction of homo-

384

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

Roche et al. logous and heterologous dihydrolipoamide dehydrogenase. Biochem J 319: 109–116, 1996. JM Jilka, M Rahmatullah, M Kazemi, TE Roche. Properties of a newly characterized protein of the bovine kidney pyruvate dehydrogenase complex. J Biol Chem 261:1858–1867, 1986. RH Behal, KS Browning, TB Hall, LJ Reed. Cloning and nucleotide sequence of the gee for protein X from Saccharomyces cerevisiae. Proc Natl Acad Sci USA 86:8732–8736, 1989. RN Perham. Swinging arms and swinging domains in multifunctional enzymes: catalytic machines for multistep reactions. Ann Rev Biochem 69:961–1004, 2000. F Hucho, DD Randall, TE Roche, MW Burgett, JW Pelley, LJ Reed. a-Keto acid dehydrogenase complexes. XVII. Kinetic and regulatory properties of pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase from bovine kidney and heart. Arch Biochem Biophys 151:328–340, 1972. FH Pettit, TE Roche, LJ Reed. Function of calcium ions in pyruvate dehydrogenase phosphatase activity. Biochem Biophys Res Commun 49:563– 571, 1972. RL Cate, TE Roche. A unifying mechanism for stimulation of mammalian pyruvate dehydrogenase kinase activity by NADH, dihydrolipoamide, acetylCoA, or pyruvate. J Biol Chem 253:496–503, 1978. LR Stepp, FH Pettit, SJ Yeaman, LJ Reed. Puriﬁcation and properties of pyruvate dehydrogenase kinase from bovine kidney. J Biol Chem 258:9454– 9458, 1983. L Li, GA Radke, K Ono, TE Roche. Additional binding sites for the pyruvate dehydrogenase kinase but not for protein X in the assembled core of the mammalian pyruvate dehydrogenase complex. Binding region for the kinase. Arch Biochem Biophys 296:497–504, 1992. GA Radke, K Ono, S Ravindran, TE Roche. Critical role of a lipoyl cofactor of the dihydrolipoyl acetyltransferase in the binding and enhanced function of the pyruvate dehydrogenase kinase. Biochem Biophys Res Comm 190:982–991, 1993. K Ono, GA Radke, TE Roche, M Rahmatullah. Partial activation of the pyruvate dehydrogenase kinase by the lipoyl-domain region of E2 and interchange of the kinase between lipoyl-domain regions. J Biol Chem 268: 26135–26143, 1993. S Liu, JC Baker, TE Roche. Binding of the pyruvate dehydrogenase kinase to recombinant constructs containing the inner lipoyl domain of the dihydrolipoyl acetyltransferase component. J Biol Chem 270:793–800, 1995. S Ravindran, GA Radke, TE Roche. Lipoyl domain–based mechanism for integrated feedback control of pyruvate dehydrogenase complex by enhancement of pyruvate dehydrogenase kinase activity. J Biol Chem 271:653–662, 1966. JC Baker, X Yan, T Peng, SA Kasten, TE Roche. Marked diﬀerences between two isoforms of human pyruvate dehydrogenase kinase. J Biol Chem 275: 15773–15781, 2000.

E2-Facilitated Kinase/Phosphatase Function

385

57. D Yang, X Gong, A Yakhnin, TE Roche. Requirements for the adaptor protein role of dihydrolipoyl acetyltransferase in the upregulated function of the pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase. J Biol Chem 273:14130–14137, 1998. 58. G Chen, L Wang, S Liu, C Chang, TE Roche. Activated function of the pyruvate dehydrogenase phosphatase through Ca2+-facilitated binding to the inner lipoyl domain of the dihydrolipoyl acetyltransferase. J Biol Chem 271: 28064–28070, 1996. 59. C-Y Maeng, MA Yazdi, X-D Niu, HY Lee, LJ Reed. Expression, puriﬁcation, and characterization of the dihydrolipoamide dehydrogenase–binding protein of the pyruvate dehydrogenase complex from Saccharomyces cerevisiae. Biochemistry 33:13801–13807, 1994. 60. JK Stoops, TS Baker, JP Schroeter, SJ Kolodziej, X Niu, LJ Reed. On the unique structural organization of the Saccharomyces cerevisiae pyruvate dehydrogenase complex. J Biol Chem 267:24769–24775, 1992. 61. RH Behal, MS DeBuysere, B Demeler, JC Hansen, MS Olson. Pyruvate dehydrogenase multienzyme complex. Characterization of assembly intermediates by sedimentation velocity analysis. J Biol Chem 269:31372–31377, 1994. 62. RG McCartney, SJ Sanderson, JG Lindsay. Refolding and reconstitution studies on the trans acetylase-protein X (E2/X) subcomplex of mammalian pyruvate dehydrogenase complex: evidence for speciﬁc binding of the dihydrolipoamide dehydrogenase component to sites on reassembled E2. Biochemistry 36:6819–6826, 1997. 63. SJ Sanderson, C Miller, JG Lindsay. Stoichiometry, organization and catalytic function of protein X of the pyruvate dehydrogenase complex from bovine heart. Eur J Biochem 236:68–77, 1996. 64. ZH Zhou, W Liao, RH Cheng, JE Lawson, DB McCarthy, LJ Reed, JK Stoops. Direct evidence for the size and conformational variability of the pyruvate dehydrogenase complex revealed by three-dimensional electron microscopy. The ‘‘breathing core’’ and its functional relationship to protein dynamics. J Biol Chem 276:21704–21713, 2001. 65. T Izard, A Aevarsson, MD Allen, AH Westphal, RN Perham, A de Kok, WG Hol. Principles of quasi-equivalence and Euclidean geometry govern the assembly of cubic and dodecahedron cores of pyruvate dehydrogenase complexes. Proc Natl Acad Sci USA 96:240–1245, 1999. 66. A Tuganova, I Boulatnikov, KM Popov. Interaction between the individual isoenzymes of pyruvate dehydrogenase kinase and the inner lipoyl-bearing domain of transacetylase component of pyruvate dehydrogenase complex. Biochem J 366:129–136, 2002. 67. T Wagenknecht, R Grassucci, GA Radke, TE Roche. Cryoelectron microscopy of mammalian pyruvate dehydrogenase complex. J Biol Chem 266:24650– 24656, 1991. 68. TE Roche, SL Powers-Greenwood, W Shi, W Zhang, SZ Ren, ED Roche, DJ Cox, CM Sorensen. Sizing of the bovine pyruvate dehydrogenase complex and

386

69.

70.

71.

72. 73.

74.

75.

76.

77.

78.

79.

Roche et al. the dihydrolipoyl transacetylase core by quasielastic light scattering. Biochemistry 32:5629–5637, 1993. TE Roche, RL Cate. Evidence for lipoic acid–mediated NADH and acetyl-CoA stimulation of liver and kidney pyruvate dehydrogenase kinase. Biochem Biophys Res Commun 72:1375–1383, 1976. RL Cate, TE Roche. Function and regulation of mammalian pyruvate dehydrogenase complex: acetylation, interlipoyl acetyl transfer, and migration of the pyruvate dehydrogenase component. J Biol Chem 254:1659–1665, 1979. M Rahmatullah, TE Roche. Modiﬁcation of bovine kidney pyruvate dehydrogenase kinase activity by CoA esters and their mechanism of action. J Biol Chem 260:10146–10152, 1985. KM Popov. Regulation of mammalian pyruvate dehydrogenase kinase. FEBS Lett 419:97–200, 1997. LG Korotchkina, MS Patel. Site speciﬁcity of four pyruvate dehydrogenase kinase isozymes toward the three phosphorylation sites of human pyruvate dehydrogenase. J Biol Chem 279:37223–37229, 2001. TE Roche, RL Cate. Puriﬁcation of porcine liver pyruvate dehydrogenase complex and characterization of its catalytic and regulatory properties. Arch Biochem Biophys 183:664–677, 1977. LG Korotchkina, MS Patel. Site speciﬁcity of four pyruvate dehydrogenase kinase isoenzymes toward the three phosphorylation sites of human pyruvate dehydrogenase. J Biol Chem 276:37223–37229, 2001. E Kolobova, A Tuganova, I Boulatnikov, KM Popov. Regulation of pyruvate dehydrogenase activity through phosphorylation at multiple sites. Biochem J 358:69–77, 2001. ML Pratt, JF Maher, TE Roche. Puriﬁcation of bovine kidney and heart pyruvate dehydrogenase phosphatase on sepharose derivatized with the pyruvate dehydrogenase complex. Eur J Biochem 125:349–355, 1982. A Turkan, X Gong, T Peng, TE Roche. Structural requirements within the lipoyl domain for the Ca2+-dependent binding and activation of pyruvate dehydrogenase phosphatase isoform 1 or its catalytic subunit. J Biol Chem 277:14976–14985, 2002. X Gong, T Peng, A Yakhnin, M Zolkiewski, J Quinn, SJ Yeaman, TE Roche. Speciﬁcity determinants for the pyruvate dehydrogenase component reaction mapped with mutated and prosthetic group–modiﬁed lipoyl domains. J Biol Chem 275:13645–13653, 2000.

22 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 University of Sheﬃeld, Sheﬃeld, England

I. INTRODUCTION In former years, studies with randomly isolated mutants having identiﬁable nutritional phenotypes made invaluable contributions to the elucidation of bacterial metabolic pathways. Now, using a combination of site-directed mutagenesis and chromosomal gene replacement, it is possible to create mutants having precisely deﬁned genetic defects but a wide range of unpredictable phenotypes. In this chapter, such ‘‘designer mutants’’ have been used to investigate the physiological consequences of altering the structure or expression of pyruvate oxidizing enzymes in E. coli. Escherichia coli uses three diﬀerent mechanisms for oxidizing pyruvate, depending on the growth conditions. First, there is the pyruvate dehydrogenase complex (PDHC), which is essential for catalyzing the NAD-linked oxidative decarboxylation of pyruvate to acetyl-CoA and CO2 and hence to funnel glycolytic carbon into the aerobic citric acid cycle. Synthesis of the PDH complex is induced by pyruvate, but its activity is inhibited under an387

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aerobic conditions by the inhibitory eﬀects of NADH on the lipoamide dehydrogenase (E3/LpdA) subunit. The PDH complex is expressed from the pdh operon (pdhR-aceEF-lpdA), consisting of a regulatory gene encoding a powerful repressor (PdhR) that blocks transcription from the pdh promoter, except in the presence of pyruvate, and three genes encoding enzymatic subunits, as shown in Figure 1 (1). All four genes are expressed from a single transcript initiating at the pdh promoter. There is also an independent lpd promoter that generates a short transcript to provide additional E3 subunits for the analogous 2-oxoglutarate dehydrogenase (ODH) complex (Fig. 1). The sucAB genes encoding the speciﬁc E1o and E2o subunits of the ODH complex are embedded in the large sdh-suc operon (sdhCDABsucABCD) that expresses three citric-acid-cycle enzymes from the sdh promoter (Refs. 2–4; Fig. 1). The sdh promoter is severely repressed under anaerobic conditions by an anaerobic repressor (ArcA) and also by excess glucose, just like the independent lpd promoter, and this provides a mecha-

Figure 1 Organization of the pdh and sdh-suc operons, encoding components of the PDH and ODH complexes of E. coli. The major promoters and transcripts (unbroken horizontal arrows) and their repression by pyruvate or by anaerobiosis (via ArcA) and excess glucose are indicated. The nucleotide substitutions conferring NADP dependence are located in the region of the black vertical bar in the lpdA gene, and the hatched region denotes the BstXI-Csp451 fragment replaced in lpdP mutants. The domain structures of the lipoate-acyltransferase subunits (E2p and E2o), including the lipoyl domains (o), are also shown.

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nism for coregulating the synthesis of E3 with the E1o and E2o subunits for assembly into the ODH complex (4). Second, pyruvate is converted to acetyl-CoA and formate by pyruvate formate-lyase (PFL), a free-radical enzyme that is induced and activated by anaerobiosis and an enzyme that is essential for acetate-independent anaerobic growth (5). Finally, the third mechanism is mediated by pyruvate oxidase (PoxB), a nonessential and seemingly wasteful membrane-associated enzyme that is induced as cultures approach stationary phase (6). It catalyzes the FAD-linked oxidation of pyruvate directly to acetate and CO2, rather than generating acetyl-phosphate like the pyruvate oxidases of lactic acid bacteria. This chapter describes the properties of mutants constructed in order to study the consequences of: (1) changing the coenzyme speciﬁcities of the PDH and ODH complexes from NAD to NADP via their shared lipoamide dehydrogenase (E3/LpdA) subunits; (2) modifying the domain organization of the lipoate acetyltransferase (E2p) subunit of the PDH complex; (3) restructuring the aceF-lpdA intergenic region of the pdh operon; and (4) up-regulating and inactivating pyruvate oxidase (PoxB). II. EFFECTS OF CHANGING THE COENZYME SPECIFICITIES OF THE PDH AND ODH COMPLEXES FROM NAD TO NADP The coenzyme speciﬁcity of lipoamide dehydrogenase (E3/LpdA) has previously been converted from NAD to NADP by substuting seven residues in its NAD-binding domain with those at the same positions in the NADPdependent glutathione reductase, see later (7). There are six essential substitutions, a single G185A substitution in the dinucleotide-binding fold and ﬁve alterations that create a positively charged nest to accommodate the 2’phosphate of NADP, and a nonessential G189A substitution. Together these changes generate a variant of LpdA, here designated LpdP, that uses NADP in preference to NAD: LpdA LpdP Gsr

180

GGGILGLEMGTVYHALGSQIDVVEMFDQVIPA GGGILALEMATVYHALGSQIDVVVRKHQVIRA 174 GAGYIAVELAGVINGLGAKTHLFVRKHAPLRS

180

A multistep strategy was accordingly devised to replace the chromosomal lpdA gene of E. coli by the lpdP gene from an LpdP expression plasmid, p116/117, kindly provided by Professor R. N. Perham (7). The plasmids used in this procedure and some of the resulting strains are illustrated in Figure 2. The ﬁrst steps involved constructing pGS954 (aceVF-lpdA : : kanR-acnB V) and the LpdA-null strain (JRG3503, lpdA : : kanR), in which a 661-bp segment of

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Figure 2 Changing the coenzyme speciﬁcity of the E3 (LpdA) subunit of the PDH and ODH complexes of E. coli to NADP. The pdh operons of plasmids and the corresponding strains are shown. The cloning-and-replacement strategy is described in the text. The lpdP gene encoding the NADP-dependent enzyme is hatched, and the central band denotes the cluster of the seven mutations. Vector DNA is denoted by thick lines, and restriction sites are abbreviated thus: B, Bcl I; Ba, BamHI; Bs, BstXI; Cs, Csp451; E, EcoRI: H, HindIII; Hc, HincII; Rv, EcoRV; Sp, SphI; Xb, XbaI; Xo, XhoI.

the lpdA gene (including the region altered in lpdP) was replaced by a kanR cassette ﬂanked by hybrid BclI/BamHI and HincII/EcoRV sites. A comparable lpdP multicopy plasmid (pGS957, aceVF-lpdP-acnBV), also containing suﬃcient ﬂanking DNA to mediate recombination with the host chromosome, was constructed by replacing most of the lpdA region of pGS954 by a unique 1.17-kb BstXI-Csp451 fragment from the lpdP coding region of pKK116/117 (Figs. 1 and 2). Construction of the lpdP mutant was then achieved nonselectively (i.e., without making any assumptions about the resulting phenotype) via in vivo chromosomal replacement of the lpdA : : kanR region of JRG3503 by the corresponding lpdP region of a thermosensitive replicon (pGS1046) analogous to pGS957 (Fig. 2). Enzymological studies with 15 independently generated lpdP mutants (represented here by JRG3756 and JRG3757) showed that the activity ratios for NADP versus NAD for E3 (Lpd) and the PDH and ODH complexes were increased approximately 105-fold relative to W3110 (lpdA+), the isogenic parental strain (Table 1). A similar switch in the activity ratios was also ob-

102 0.8 0.2 0.3 <0.01

W3110 lpdA+ JRG3756 lpdP JRG3757 lpdP JRG3503(pGS957) lpd(lpdP) JRG3503 lpdA :: kanR 0.2 114 76 130 0.1

NADP 0.002 142 380 433 —

NADP NAD 27 0.4 0.3 0.3 0.06

NAD <0.01 30 29 35 0.03

NADP

PDHC

<0.0003 75 97 98 —

NADP NAD

4.6 0.01 0.01 0.02 <0.01

NAD

<0.01 1.3 3.4 2.8 <0.01

NADP

ODHC

<0.002 130 340 140 —

NADP NAD

Cell-free extracts of the parental strain (W3110), representative lpdP mutants, an E3 (Lpd) null strain (JRG3503), and a derivative of the latter containing the multicopy lpdP plasmid (pGS957), were assayed for E3 (Lpd), PDHC, and ODHC activities with NAD and NADP as coenzymes. Average speciﬁc activities (Amol coenzyme reduced/mg protein/h) are quoted for three independent cultures.

NAD

E3 (Lpd)

Enzymological Analysis of Strains Having Altered PDH and ODH Complex Coenzyme Speciﬁcities

Strain

Table 1

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served with transformants of the lpdA : : kanR mutant (JRG3503) containing multicopy pGS957 (lpdP) (see Table 1) or cointegrant derivatives of the same strain containing a single copy of the thermosensitive replicon pGS1046 (not shown). Nucleotide sequence analyses with PCR-ampliﬁed DNA conﬁrmed the enzymological evidence that the desired lpdA gene replacement had been achieved in the lpdP mutants. Growth tests using a variety of substrates and conditions later showed that the lpdP mutants behave like slightly leaky lpdA mutants. Indeed, they were almost as impaired as the E3 (LpdA) null strain (JRG3503) in glucose minimal media (Fig. 3a). They resembled lpdA mutants in failing to respond to acetate alone, but they grew slightly better with succinate and a dual supplement of acetate and succinate (which allows good growth, but not as good as the wild type). The lpdP multicopy plasmid (pGS957) failed to complement the nutritional lesion of the lpdA mutant, unlike the lpdA plasmid, pGS956 (Fig. 3b). Small enhancements were observed in unsupplemented and single supplemented media, but growth with the dual supplement was actually im-

Figure 3 Growth of strains having NADP-dependent PDH and ODH complexes. (a) Growth of a representative lpdP mutant (JRG3756; unbroken line) compared to the lpdA mutant (JRG3503; dashed line) and the lpd+ strain (W3110; *), in glucose minimal medium with: a, acetate: s, succinate; a+s, acetate and succinate; or -, no supplement. (b) Complementation by the lpdP plasmid (pGS957; bold lines with symbols) compared to the lpdA plasmid (pGS956; bold unbroken line without symbols) and the untransformed lpdA host (JRG3503; dashed line) in glucose minimal medium with supplements as in panel (a).

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paired by mutiple copies of the lpdP gene. Thus it was concluded that NADP cannot be used in vivo as a redox cofactor for the oxidation of pyruvate and 2-oxoglutarate. Attempts were subsequently made to select derivatives of the lpdP strains that could use the NADP-dependent complexes, e.g., by altering the expression of transhydrogenase or by invoking some other recycling pathway from NADPH to NAD possibly via a thiol reductase. This yielded partial revertants of the types isolated previously, Sin+, which conserve succinate by inactivating succinate dehydrogenase, and Ain+, which that provide acetate by up-regulating pyruvate oxidase (8), but no respiration-proﬁcient nutritionally independent revertants were recovered. Clearly, NADP cannot be used as the redox cofactor for pyruvate and 2-oxoglutarate catabolism, indicating that the reoxidation of NADPH is not readily achieved or easily induced by mutation in E. coli. If the two nucleotide pools are eﬀectively compartmentalized by the kinetic properties of relevant enzymes that maintain very high NAD+/NADH ratios and very low NADP+/NADPH ratios, as in the mammalian cytosol (9), there might be insuﬃcient NADP to mediate pyruvate and 2-oxoglutarate metabolism at the rate needed to support growth of the lpdP strain. III. EFFECTS OF RESTRUCTURING THE aceE-lpdA REGION OF THE pdh OPERON ON THE SUBUNIT STOICHIOMETRIES OF OVEREXPRESSED PDH COMPLEXES The PDH complex can be ampliﬁed to 30% of soluble cell protein from expression plasmids, but after puriﬁcation such complexes are invariably deﬁcient in E1p and overloaded with E3 (10). The subunit stoichiometries (E1p: E2p : E3) are approximately 0.4 : 1.0 : 1.2, rather than the ‘‘textbook’’ ratio of 1.0 : 1.0 : 0.5, and the speciﬁc activities are about 40% lower than those of complexes puriﬁed from unampliﬁed sources, probably reﬂecting their lower E1p contents. The ampliﬁed cultures also contained a large excess of unbound E3 subunits, suggesting that the independent lpd promoter is deregulated in multicopy situations. Attempts were therefore made to improve the subunit stoichiometries and hence to increase the activity of ampliﬁed complexes by deleting the independent lpd promoter so that E3 expression is initiated, together with the E1p and E2p subunits, solely from the upstream pdh promoter (Fig. 1). In the pdh operon, the aceF and lpdA genes are separated by a large and complex intergenic region of 324 bp (Fig. 4). This contains a stem-loop structure resembling a transcription terminator (Tace), which might serve to halve lpdA expression relative to the aceEF genes (consistent with the 1.0:1.0:0.5 subunit stoichiometries), but it is more probably a processing site

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Figure 4 Eﬀects of deletions in the aceF-lpdA intergenic region. Features of the 324bp intergenic region are illustrated for ptac-85 expression plasmids and the corresponding bacterial chromosomes: wild-type parent (pGS501, W3110); (DP, a 248-bp lpd promoter deletion (pGS580, JRG4035); and (DTP, a 298-bp transcript processing site and lpd promoter deletion (pGS579, JRG4034). Transcription start sites are denoted by arrows and the ArcA binding-site palindrome and the potential stem-loop of the intergenic repeat unit are also indicated. The activities of PDHC and E3 (LpdA) in cell-free extracts of plasmid-containing cultures treated with and without IPTG are recorded (Amol NAD+ reduced/mg protein/min), as are the subunit stoichiometries and speciﬁc activities of the overexpressed and puriﬁed complexes.

for the read-through transcript (4). Then there is an ArcA-protected region containing an ArcA binding-site palindrome ﬂanked by the -35 and -10 hexamers of the independent lpd promoter. And ﬁnally, between the lpd transcript start site and lpdA coding region is a 127-bp intergenic repeat unit (IRU) or enterobacterial repetitive intergenic consensus (ERIC) sequence (Fig. 4). This sequence forms part of every aceEF-lpdA and lpdA transcript; but like all of the 19 such sequences in the E. coli chromosome, it has no known function (11). Two deletions were engineered. One, of 248 bp (DP), removes the lpd promoter and the IRU but not the processing site (Tace) and the other, of 298 bp (DTP), removes the entire intergenic region, except for the aceF stop codon and the 24-bp segment between a XhoI site and the lpdA gene (Fig. 4). The deletions were constructed in a Ptac-aceEF-lpdA expression plasmid, pGS501 (10), and transferred to the chromosomal pdh operons of the parental strain (W3110). As expected, synthesis of PDHC was entirely dependent on induction by IPTG in all cases (Fig. 4). However, the E3 (LpdA) activities indicated that contrary to expectations, lpdA expression remained independent of IPTG with the smaller deletion (DP); but as predicted, it did become almost entirely IPTG dependent with the larger deletion (DTP). This anomaly

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was later explained by transcript analysis and start-site analysis with the corresponding bacterial strains, which showed that a new independent lpd promoter becomes active in the residual intergenic region of the DP construct (as indicated by the arrowhead in Fig. 4), whereas for the DTP construct lpdA expression depends almost entirely on the upstream tac promoter. Unfortunately, neither deletion improved the subunit stoichiometries or the speciﬁc activities of the complexes puriﬁed from the corresponding ampliﬁed sources. An excess of unbound E3 was still produced in both cases, presumably from the new independent lpd promoter with the DP expression plasmid and for unknown reasons with the DTP plasmid, where equal relative amounts of all three subunits would have been expected if the three adjacent genes were transcribed and translated without modulation. An excess of unbound E3 expressed from a deregulated or independent lpd promoter in multicopy situations might be responsible for displacing E1 subunits during assembly of the PDH complex. However, it also seems likely that the subunit stoichiometries of the complex are normally controlled by other more subtle features modulating transcription and translation. Growth tests with the DP and DTP mutant strains conﬁrmed that they synthesize suﬃcient E3 for ODHC assembly and growth in unsupplemented media containing glucose, succinate, and acetate as sole carbon and energy sources. Quantitative growth tests further conﬁrmed that growth on glucose is unimpaired and likewise for growth on acetate, which is somewhat surprising for the DTP strain, where lpdA expression depends entirely on the pdh promoter, which is repressed because PDHC is not required for growth on acetate. However, growth on succinate was increasingly impaired in the DP and DTP mutants, in a way that could be compensated by added acetate. So it would appear that removing the lpd promoter does aﬀect the use of succinate, i.e., where the organism is most dependent on a supply of E3 subunits for both the PDHC and ODHC. IV. THREE LIPOYL DOMAINS PER ACETYLTRANSFERASE CHAIN IS BEST Ever since the aceF and sucB genes revealed the primary structures of the E2 subunits it has been asked why the lipoate-acetyltransferase (E2p) has three tandemly repeated lipoyl domains per E2p chain, whereas the succinyltransferase functions with only one (Fig. 1). In seeking an answer, genetically engineered multilip PDH complexes containing from one to eight lipoyl domains per chain were constructed, one series (+/+n) containing lipoyl domains that are all lipoylatable, and another series (+/n) containing just one lipoylatable domain at the N-terminus, the rest being blocked by a Lys!Gln substitution at the lipoylation site (12,13). However, the speciﬁc

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activities of both types of complex declined with increasing numbers of lipoyl domains rather than showing a preference for three (Fig. 5a). Indeed, the highest activities were observed with only one lipoyl domain per E2p chain, nor was any advantage conferred by the presence of extra lipoyl cofactors, because both types of 3lip complex (+++ and +– –) were equally active. The presence of extra domains was inhibitory, but even the 8lip complex having a single lipoyl cofactor at the N-terminal extremity retained almost 20% of wild-type activity, indicating that its remote cofactors have the ability to access all three types of active site in the assembled complex (Fig. 5a).

Figure 5 Properties of multilip PDH complexes and isogenic strains expressing these complexes from chromosomal pdh operons. (a) Comparisons for speciﬁc activities of overexpressed and puriﬁed PDH complexes (thin lines) and maximum speciﬁc growth rates in glucose minimal medium (thick lines) versus number of lipoyl domains per E2p chain. Data for two series of complexes, containing entirely lipoylatable domains (+/+n) or single N-terminal lipoylatable domains connected to increasing numbers of nonlipoylatable domains (+/n), are each related to the wild type (W3110; 3lip +++), where 100% =12 Amol/mg protein/min or 0.7 h1 (13). (b) Pairwise competition between equal numbers of genetically marked wild-type and unmarked isogenic multilip mutants growing in glucose-limited chemostat cultures. The numbers of mutants, expressed as a fraction (%) of total viable bacteria, is plotted logarithmically against the number of generations at the time of sampling (12,13). A control experiment containing marked and unmarked wild types (3lip, +++) is also shown.

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Enzymological studies of this type not only indicated that the wild-type complex possesses superﬂuous lipoyl domains and lipoyl cofactors, but selective deletions within the Ala+Pro-rich lipoyl linkers of the 1lip complex further indicated that the linkers are approximately 40% longer than needed: 12 of the 32 residues could be deleted without loss of activity (14). It was also found that the 25% loss of activity caused by deleting all but 14 residues was more than compensated by inserting a 14-residue poly-Pro linker, which improved activity to 180% of wild type (15). The enhanced activity conferred by the poly-Pro linker was attributed to its greater reach and lower ﬂexibility compared to an Ala+Pro-rich linker, which needs to be longer to function optimally. Because the enzymological approach oﬀered no explanation for the retention of three lipoyl domains per E2p subunit, an alternative ‘‘whole organism’’ strategy was adopted. This involved replacing the chromosomal pdh operons by those encoding each of the multilip complexes, to see whether the physiological properties of growing cultures might signal a preference for the 3lip complex. Using intact genetically engineered organisms avoids complications arising from variations in the activity, purity, subunit stoichiometry, and degree of lipoylation that might occur between diﬀerent preparations of the overexpressed PDH complexes. Indeed, full lipoylation could only be guaranteed for complexes expressed from single-copy chromosomal operons compared to overexpressed complexes. So if it is assumed that bacteria growing under identical conditions express and assemble the complexes in identical fashion, the ‘‘whole organism’’ strategy allows the bacteria to do most of the work and report the eﬃciencies of the complexes in simple physiological terms. The growth rates measured in controlled batch culture clearly indicate that a 3lip complex is better than a 1lip, 2lip, or >3lip complex (Fig. 5a; Ref. 13). It is also very signiﬁcant that the same optimal growth rate is supported by both types of 3lip complex, +++ and +– – (Fig. 5a). Furthermore, detailed physiological analyses of both 3lip strains failed to reveal any diﬀerences in growth rate (Amax), molar growth yield, or carbon conversion eﬃciency. This was true for chemostat cultures growing with limiting substrates such as glucose, pyruvate, lactate, and succinate, which require PDHC for their metabolism. Strains expressing the 2lip complexes (++ and +) exhibited identical but suboptimal growth rates (Fig. 5a), and the growth rates of strains having more than three lipoyl domains per E2p chain declined as the number increased, indicating that extra domains impair function. The growth rates of the +/n series were consistently better than those of the +/+n series. This suggests that extra lipoylated domains may be more inhibitory than nonlipoylated domains, or, alternatively, if the extra lipoyl domains of the +/+n series are not fully lipoylated some complexes

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would have unlipoylated domains at or near the N-terminal ends, an organization that was earlier found to impair function in comparisons between – –+ and +++ complexes (16). A detailed physiological analysis of two 1lip strains containing short (14-residue) lipoyl linkers of diﬀerent compositions, Ala+Pro (JRG4157) and poly-Pro (JRG4164), showed that all of the major growth constants are not signiﬁcantly diﬀerent from those of the parental 1lip strain (JRG2931), despite earlier observations (15) that the speciﬁc activities of the isolated PDH complexes are, respectively, lower and higher than the parental 1lip complex. A more stringent test of physiological performance is to set up glucoselimited chemostat cultures inoculated with equal numbers of mutant bacteria and a marked wild-type strain and then to follow the retention of the mutant as a fraction of the total population, over many generations. Such pair-wise competition experiments showed that the 2lip, 1lip, and 6 lip bacteria are displaced from the cultures at increasing rates (Fig. 5b). However, both types of 3lip strain, +++ and +– –, were retained at a constant fraction of 50% for at least 50 generations, indicating that they are equally eﬃcient and hence indistinguishable by physiological criteria. The physiological studies have thus provided the ﬁrst clear evidence that three is the optimal number of lipoyl domains per E2p chain in the PDH complex. Prior to this, the only support for this preference was inferred from NMR studies that showed that the 3lip complex exhibits a greater inherent mobility (linker:domain intensity ratio) than any of the other complexes (17). The other conclusions arising from the physiological studies can be summarized as follows. 1. Three lipoyl domains per E2p chain is best, not because it provides extra lipoyl cofactors, but because it optimizes the reach of the Nterminal lipoyl cofactor. Presumably, this situation has evolved by lipoyl domain and linker duplication during the evolution of the multienzyme complex into an organized arrangement of components allowing the highest eﬃciency of active-site coupling. 2. It can be inferred that in the assembled complex where the outer shell of E1p and E3 subunits and the inner core of E2p subunits deﬁne the space in which the lipoyl cofactors operate, the 3lip ‘‘domain-linker’’ arrangement optimizes the N-terminal lipoyl cofactor’s ability to reach across the space to interact with the active sites. 3. The complexes must have open, ﬂexible structures that permit a high degree of freedom for the lipoyl domains and lipoyl cofactors to visit each component enzyme. Were it otherwise, it would be

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diﬃcult to explain how 1lip-to-8lip complexes containing single lipoyl cofactors at the N-terminal extremities of the E2p subunits support growth rates >50% of the wild-type rate. 4. The PDHC-speciﬁc activities measured under substrate-saturating conditions do not truly reﬂect physiological performance in vivo. Depending on the number of lipoyl domains, these speciﬁc activities both overestimate and underestimate their performance relative to the physiological competence of the corresponding strains. Similar discrepancies were observed after changing the length and composition of the lipoyl linkers. In future it would be interesting to conduct evolutionary studies with glucose-limited cultures over many generations, to see whether superﬂuous domains are deleted from the >3lip strains to generate the optimal 3lip arrangement. Such studies might likewise show whether 1lip and 2lip strains can evolve by increasing the number of lipoyl domains to three or by changing the length or composition of the lipoyl linkers in response to prolonged nutritional limitation. Such experiments would be relatively easy to perform using a pair of speciﬁc PCR primers ﬂanking the lipoyl domain and linker regions of the pdh operon to monitor potential changes in the cultures. It might also be informative to investigate the responses of strains having suboptimal linker lengths and composition during prolonged exposure to nutritional stress. V. THE PHYSIOLOGICAL CONSEQUENCES OF REPLACING THE PDH COMPLEX BY PYRUVATE OXIDASE The pyruvate oxidase (PoxB) of E. coli has been much studied in the Hager, Gennis, and Cronan laboratories at the University of Illinois. It is a rather enigmatic TDP-dependent ﬂavoprotein that oxidizes pyruvate directly (and seemingly wastefully) to acetate, rather than to acetyl-phosphate like the Lactobacillus plantarum enzyme or to acetyl-CoA like the PDH complex. It is synthesized as cultures approach stationary phase under the control of RpoS (j38) the stationary-phase sigma factor (6). Pyruvate oxidase is not essential for normal aerobic growth, nor does it supply suﬃcient acetate to support the growth of mutants lacking the PDH complex. Recently, attempts were made to see whether PoxB could be made to replace PDHC for supporting acetate-independent growth and, if so, what physiological or energetic consequences might stem from such a replacement (18). For this purpose a series of isogenic mutants of wild-type E. coli (W3110) were constructed. As shown in Table 2, this series included: a PoxB null strain having a kanR

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Table 2 Physiological Eﬀects of Blocking the PDH Complex and PoxB Routes of Pyruvate Oxidation

Genotype

Phenotype

Enzymes in strain

Growth rate max Amax(h1)

Carbon eﬃciency (%)

Growth yield (g/mol)

W3110 JRG3456 JRG3980 JRG3445

Wild type aceE poxB+ aceE poxBc aceE Ptac-poxB+

Ace+ Ace Ace+ Ace

JRG3434 JRG3931

aceE+ poxB aceE poxB

Ace+ Ace

PDHC + PoxB PoxB PoxB — (+ acetate) PoxB (+ IPTG) PDHC —

0.69* 0.46 0.46* 0.36 0.46* 0.58* 0.36

57 41 35 — 36 40 —

93 78 70 — 85 80 —

Strain

Maximum speciﬁc growth rates were determined for controlled batch cultures of isogenic strains in glucose minimal medium supplemented with acetate (2 mM ), whether needed or not, or with IPTG (50 lM ) where indicated (*denotes that growth rates were unaﬀected by omitting acetate). Other parameters were determined for glucose-limited chemostat cultures maintained at steady states over a range of growth (dilution) rates in minimal media without supplements of acetate or IPTG where possible, except for strains having Ace phenotypes. Source: Ref. 18.

insertion in the poxB coding region (JRG3434); a PDH-E1p null strain in which part of the aceE gene had been replaced by a camR cassette (JRG3456); and further derivatives of the PDH-E1p null strain containing the inactivated poxB gene (JRG3931) with either, a poxBc mutation selected for constitutive rather than stationary-phase dependent expression of PoxB (JRG3980), or a poxB gene that is expressed from the IPTG-inducible tac promoter (JRG3445). Enzymological tests showed that PoxB activities are maintained throughout the growth cycle at twice the normal stationary-phase level in the poxBc strain and are ampliﬁed 100-fold in the Ptac-poxB strain when induced with IPTG (18). Aerobic growth tests in glucose minimal media conﬁrmed that the parental and PoxB-deﬁcient strains are acetate independent (Ace+ phenotype) and that the PDHC-deﬁcient strain requires acetate (Ace). However, the PDHC-deﬁcient poxBc strain was found to be Ace+, and the PDHC-deﬁcient Ptac-poxB strain had an Ace phenotype that became Ace+ when grown in the presence of IPTG (Table 2). Hence it is clear that PoxB can replace PDHC and permit good growth and colony formation, if it is expressed continuously throughout the growth cycle from a constitutive promoter or from the externally regulated Ptac promoter rather than the natural promoter. Tests

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using chemostat cultures have further shown that the naturally expressed PoxB can indeed support growth of a PDHC-deﬁcient mutant in unsupplemented medium, but only at low growth rates (V0.05 h1) as compared to rates of up to 0.25 h1 (the highest tested) observed with the poxBc and PtacpoxB strains (18). In order to detect any physiological or energetic consequences stemming from the use of PoxB instead of PDHC, quantitative growth tests were performed with controlled batch cultures and also with glucose-limited chemostat cultures maintained at several diﬀerent growth rates. These showed that in the absence of a PDH complex, the maximum speciﬁc growth rate is a third lower, the carbon conversion eﬃciency (i.e., the conversion of substrate to biomass) is reduced by the same amount, and the growth yield per mole of substrate consumed is also lower, in all of the strains relying on PoxB, however it is expressed (Table 2). So compared to the wild type, more substrate has to be consumed to achieve a given yield when there is no PDHC, and PoxB provides the only route for funneling pyruvate carbon into the citric acid cycle. This is undoubtedly due to pyruvate oxidation via acetate being less eﬃcient than the direct route to acetyl-CoA; see Figure 6. First, the reducing equivalents are recovered at the less favorable FADH2 level with PoxB, rather than as NADH. Then, in order to convert the acetate product to acetyl-CoA, ATP has to be consumed by acetyl-CoA synthetase (ACS), which is induced 13- to 20-fold in the PDHC-deﬁcient cultures relative to the PoxBdeﬁcient strain (18). It is therefore concluded that the PoxB+ACS route can replace the PDHC route, but less eﬃciently.

Figure 6 Two routes for the aerobic conversion of pyruvate to acetyl-CoA and CO2. The PDH complex catalyzes the NAD-dependent oxidative decarboxylation of pyruvate directly to acetyl-CoA, whereas the pyruvate oxidase (PoxB)-mediated route involves the FAD-dependent conversion of pyruvate to acetate followed by the ATP-dependent synthesis of acetyl-CoA by acetyl-CoA synthetase (ACS).

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VI. PYRUVATE OXIDASE CONTRIBUTES TO THE GROWTH EFFICIENCY OF E. coli The greater eﬃciency of the PDHC-mediated route of pyruvate oxidation relative to the PoxB+ACS route, raises questions concerning why E. coli retains PoxB and the ineﬃcient PoxB+ACS route. Presumably the organism derives some beneﬁt from retaining and expressing PoxB, albeit primarily as cultures enter the stationary phase. A detailed physiological analysis of the PoxB-null strain (JRG3434) has indeed shown that the growth rate (Amax), the carbon conversion eﬃciency, and the molar growth yield are all lower than the parental values (Table 2). Likewise, for strains lacking the PDH complex, the growth rate is signiﬁcantly lowered (from 0.46 to 0.36 h1) when PoxB is also inactivated (JRG3931) or when the Ptac-poxB gene is not induced (JRG3445). These observations clearly show that PoxB makes a signiﬁcant contribution to the aerobic growth eﬃciency of E. coli. It is not known how this is achieved, but it has been suggested that the production of acetate rather than acetyl-CoA may conserve the CoA pool for other metabolic functions, such as the conversion of 2-oxoglutarate to succinyl-CoA, thus maintaining citric acid cycle activity (18). Alternatively, the exclusive use of the PDHC route could have a detrimental eﬀect on cellular redox balance by lowering the NAD+/NADH ratio, and this might be avoided by using the PoxB route to generate reducing equivalents at the ﬂavin level. The existence of two pyruvate oxidation routes parallels that of the two diﬀerentially regulated NADH dehydrogenases: NdhI, the proton-translocating complex, and NdhII, the non-proton-translocating ﬂavoprotein (19–21). NdhI is used during carbon limitation, low aeration, and late in the growth cycle, when energy conservation is most needed, whereas NdhII (which is equivalent to PoxB) is used preferentially during aerobic respiration. It would thus appear that there are conditions where energy conservation per se is not a major factor in deﬁning growth eﬃciency, and it is concluded that there must be some advantage to be gained from oxidizing at least a fraction of glycolytic pyruvate via the wasteful route.

VII. CONCLUSIONS Advances in molecular genetics have provided methods for inducing mutations and creating mutant organisms by design rather than at random (reverse genetics). These mutants can now be widely used, not only to investigate the structure–function relationships of proteins but also to study the physiological consequences of lesions that have more subtle eﬀects on growth and viability than those isolated previously that had to have clear-cut growth

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defects and selectable phenotypes. Examples of both types of applications have been explored here in the context of the PDH complex and pyruvate metabolism in E. coli. Changing the coenzyme speciﬁcity of the E3 component of the PDH and ODH complexes, from NAD to NADP, revives interest in the metabolic compartmentalization of these redox cofactors from an earlier metabolic era and oﬀers new ways of approaching the subject. Attempts to improve the subunit stoichiometries of overexpressed PDH complexes by removing the independent lpd promoter with part or most of the aceF-lpdA intergenic region of the pdh operon failed. This indicates that the abnormal subunit stoichiometry is not due simply to transcription-dependent overproduction of E3 subunits but to other factors. It was also interesting to ﬁnd that creating a monocistronic pdh operon in the DTP strain induced only mild symptoms of E3 (LpdA)/ODHC deﬁciency. However, this deﬁciency was most manifest during growth on succinate, where the demand for both PDHC and ODHC is greatest, and this observation does at least show that there are conditions that beneﬁt from the presence of an independently regulated lpd promoter. It might have been interesting to make a detailed physiological analysis of the DP and DTP strains, including competition experiments with wild type, in order to quantitate the extents of their debilitation. Studies with the multilip strains were particularly important for providing the ﬁrst clear evidence that the wild-type PDHC having three lipoyl domains per E2p is the most eﬃcient under physiological conditions, despite conﬂicting enzymological evidence. However, the preference for three domains is not due to the need for extra cofactors, but because they optimize the reach of the N-terminal lipoyl cofactor. Presumably this preference reﬂects the distance between the E2p core and the three types of active site visited by the lipoyl cofactor. The fact that the extended reach of the critical cofactor seems to have been achieved by duplicating an existing structural element, rather than recruiting an unlipoylated structure or extended linker, supports the well-established evolutionary principle of incremental modiﬁcation of existing material. Some of the engineered strains could in future be used in evolutionary experiments to investigate how the numbers of domains and linker compositions improve when strains having suboptimal structures are subjected to prolonged nutritional stress. Finally, some of the mysteries concerning the seemingly wasteful pyruvate oxidase have been resolved by showing that when expressed continuously, PoxB can be used to support the growth of E. coli in the absence of PDHC, albeit less eﬃciently. Even more surprising, the physiological analysis showed that PoxB actually contributes to the growth eﬃciency under normal conditions (i.e., in the presence of a functional PDHC). It explains why PoxB has been retained by E. coli but now poses questions about how the greater

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eﬃciency is achieved. Again, some of the newly constructed strains and plasmids could be used very proﬁtably in future physiological and protein structure–function studies. In particular, the double mutant lacking PDH-E1p and PoxB (JRG4077, aceE :: camR poxB :: kanR) has several uses. It provides a null background, allowing each enzyme to be assayed without interference from the pyruvate dehydrogenase/oxidase activity of the other. It would serve as an invaluable host for detailed molecular–genetic and structure–function analyses of the PoxB enzyme using cloned and manipulated poxB genes. The double mutant also has some biotechnological potential because although it requires acetate as an essential but titratable growth supplement, it cannot accumulate this major inhibitor of biomass production. The PDH-defective PoxB-constitutive strain (JRG3980, aceE :: camR poxBc) would also provide a very useful parental strain for selecting poxB mutants, which would acquire an Ace phenotype, and for other conventional genetic studies. ACKNOWLEDGMENTS Some of this work has been supported at diﬀerent times over several years by studentships and project grants from the Biotechnology and Biological Research Council, the Wellcome Trust, the CNPq Brazil, and the Egyptian Education Bureau. REFERENCES 1.

2. 3.

4.

5.

6.

MA Quail, JR Guest. Puriﬁcation, characterization and mode of action of PdhR: the transcriptional repressor of the pdhR-aceEF-lpd operon of Escherichia coli. Mol Microbiol 15:519–529, 1995. L Cunningham, JR Guest. Transcription and transcript processing in the sdhCDAB-sucABCD operon of Escherchia coli. Microbiol 144:213–2123, 1998. S-J Park, G Chao, RP Gunsalus. Aerobic regulation of the sucABCD genes of Escherichia coli, which encode a-ketoglutarate dehydrogenase and succinyl coenzyme A synthetase: roles of ArcA, Fnr, and the upstream sdhCDAB promoter. J Bacteriol 179:4138–4142, 1997. L Cunningham, D Georgellis, J Green, JR Guest. Co-regulation of lipoamide dehydrogenase and 2-oxoglutarate dehydrogenase synthesis in Escherichia coli: characterization of an ArcA-binding site in the lpd promoter. FEMS Microbiol Lett 169:403–408, 1998. D Kessler, J Knappe. Anaerobic dissimilation of pyruvate. In: Neidhardt FC, ed. Escherichia coli and Salmonella: Cellular and Molecular Biology. Washington, DC: ASM Press, 1996, pp 199–205. Y-Y Chang, A-Y Wang, JE Jr Cronan. Expression of Escherichia coli pyruvate oxidase (PoxB) depends on the sigma factor encoded by the rpoS (katF ) gene. Mol Microbiol 11:1019–1028, 1994.

Effects of Replacing PDH Complex of E. coli 7.

8.

9.

10.

11. 12.

13.

14.

15.

16.

17. 18.

19.

20.

21.

405

JA Bocanegra, NS Scrutton, RN Perham. Creation of an NADP-dependent pyruvate dehydrogenase multienzyme complex by protein engineering. Biochemistry 32:2737–2740, 1993. IT Creaghan, JR Guest. Suppression of the succinate requirement of lipoamide dehydrogenase mutant of Escherichia coli by mutations aﬀecting succinate dehydrogenase activity. J Gen Microbiol 102:183–194, 1977. KA Gumaa, P McLean, AL Greenbaum. Compartmentation in relation to metabolic control in liver. In: PN Campbell, F Dickens, eds. Essays in Biochemistry, Vol. 7. London: Academic Press, 1971, pp 39–86. GC Russell, RS Machado, JR Guest. Overproduction of the pyruvate dehydrogenase multienzyme complex of Escherichia coli and site-directed substitutions in the E1p and E2p subunits. Biochem J 287:611–619, 1992. FR Blattner, sixteen others. The complete genome sequence of Escherichia coli K-12. Science 277:1453–1462, 1997. E Dave´, JR Guest, MM Attwood. Metabolic engineering in Escherichia coli: lowering the lipoyl domain content of the pyruvate dehydrogenase complex adversely aﬀects the growth rate and yield. Microbiology 141:1839–1849, 1995. JR Guest, MM Attwood, RS Machado, KY Matqi, JE Shaw, SL Turner. Enzymological and physiological consequences of restructuring the lipoyl domain content of the pyruvate dehydrogenase complex of Escherichia coli. Microbiology 143:457–466, 1997. JS Miles, JR Guest, SE Radford, RN Perham. Investigation of the mechanism of active site coupling in the pyruvate dehydrogenase multienzyme complex of Escherichia coli by protein engineering. J Mol Biol 202:97–106, 1988. SL Turner, GC Russell, MP Williamson, JR Guest. Restructuring an interdomain linker in the dihydrolipoamide acetyltransferase component of the pyruvate dehydrogenase complex of Escherichia coli. Prot Eng 6:101–108, 1993. AG Allen, RN Perham, N Allison, JS Miles, JR Guest. Reductive acetylation of tandemly-repeated lipoyl domains in the pyruvate dehydrogenase multienzyme complex of Escherichia coli is random order. J Mol Biol 208:623–633, 1989. RS Machado, JR Guest, MP Williamson. Mobility in pyruvate dehydrogenases with multiple lipoyl domains. FEBS Lett 323:243–246, 1993. AM Abdel-Hamid, MM Attwood, JR Guest. Pyruvate oxidase contributes to the aerobic growth eﬃciency of Escherichia coli. Microbiology 147:1483–1498, 2001. RB Gennis, V Stewart. Respiration. In: Neidhardt FC, ed. Escherichia coli and Salmonella: Cellular and Molecular Biology. Washington, DC: ASM Press, 1996, pp 217–261. MW Calhoun, KL Oden, RB Gennis, MJ Teixeria de Mattos, OM Neijssel. Energetic eﬃciency of Escherichia coli: eﬀects of mutations in components of the aerobic respiratory chain. J Bacteriol 175:3020–3025, 1993. QH Tran, J Bongaerts, D Vlad, G Unden. Requirement for the proton-pumping NADH dehydrogenase I of Escherichia coli in respiration of NADH to fumarate and its bioenergetic implications. Eur J Biochem 244:155–160, 1997.

23 Structure and Intersubunit Information Transfer in the E. coli Pyruvate Dehydrogenase Multienzyme Complex William Furey, Palaniappa Arjunan, Andrew Brunskill, and K. Chandrasekhar Veterans Aﬀairs Medical Center and University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, U.S.A. Natalia S. Nemeria, Wen Wei, Yan Yan, Sheng Zhang, and Frank Jordan Rutgers University, Newark, New Jersey, U.S.A.

I. INTRODUCTION The Escherichia coli pyruvate dehydrogenase multienzyme complex (PDHc) catalyzes the oxidative decarboxylation of pyruvate in the following overall reaction (Scheme 1) (1): Pyruvate þ CoA þ NADþ ! Acetyl CoA þ CO2 þ NADH þ Hþ

ð1Þ

In E. coli three diﬀerent enzyme components are involved in this reaction: E1, or pyruvate dehydrogenase, utilizing TDP as a cofactor (EC1.2.4.1;E1); E2, or dihydrolipoamide transacetylase, which contains covalently bound lipoyl 407

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Scheme 1

groups (EC2.3.1.12; E2); and E3, or lipoamide dehydrogenase, containing tightly bound FAD (EC1.8.1.4; E3). The multienzyme complex performs the following series of reactions (2–4). Pyruvate þ E1 TDP Mg2þ ! E1 hydroxyethylidene TDP Mg2þ þ CO2 E1 hydroxyethylidene TDP Mg2þ þ E2 lipoamide ! E1 TDP Mg2þ þ E2 acetyldihydrolipoamide E2 acetyldihydrolipoamide þ CoA ! E2 dihydrolipoamide þ acetyl CoA E2 dihydrolipoamide þ E3 FAD ! E2 lipoamide þ E3 FADH2 E3 FADH2 þ NADþ ! E3 FAD þ NADH þ Hþ

ð2Þ ð3Þ ð4Þ ð5Þ ð6Þ

The complex consists of multiple copies of each component, with a polypeptide stoichiometry of: 24 E1, mass 99,474 Da (5): 24 E2, mass 65,959 Da (6); and 12 E3, mass of 50,554 Da (7); for a total weight of 4.57 106. The recently completed crystal structure of the E. coli pyruvate dehydrogenase complex E1 component (PDHc-E1) has enabled us to probe the role of amino acid side chains near the catalytic thiamine diphosphate (TDP).

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II. STRUCTURAL INFORMATION ON THE E. coli PDHc-E1 A. General Description and Subunit Assembly The crystal structure of the E. coli PDHc-E1 component has been determined at 1.85-A˚ resolution (8). This work represents the ﬁrst reported structural example of an E1 component from the a2 homodimeric class and also the ﬁrst example of a functional E1 of any sort with speciﬁcity for pyruvate. The structure is generally well ordered, but in the electron density maps there are three regions of weak (uninterpretable) or missing density corresponding to residues 1–55, 401–413, and 541–557. Accordingly, no detailed structural information is available for these regions. Mass spectrometric analyses on washed and dissolved crystals, however, showed that these regions are in fact present and therefore must simply be locally disordered in the crystal, rather than having been excised by proteolysis. Each PDHc-E1 subunit contains a single chain of 886 amino acids and is partitioned into three domains. The Nterminal domain is the largest of the three and contains two of the three disordered regions. Two identical subunits assemble to form a tightly packed a2 homodimer that represents the active E1 enzyme with dimensions of roughly 103 95 71 A˚. The rms deviation between corresponding Ca atoms in the dimer is 0.37 A˚, indicating virtually identical folding patterns in the two subunits. In the dimer the two subunits are related by a noncrystallographic twofold axis, and in the subunit–subunit interface two symmetrically related TDP-binding pockets form the enzyme’s active sites. Individual domain, subunit, and dimer structures are illustrated in Figure 1a and 1b, respectively. B. Cofactor Binding and Active-Site Environment As in all TDP-dependent enzymes of known structure, dual active sites are formed at the interface between subunits comprising a tightly packed dimer with conserved cofactor binding interactions involving both subunits, and the TDP cofactors are held in the V-conformation. In E. coli, PDHc-E1 conserved protein–cofactor interactions include a side-chain E571-N1V hydrogen bond, a main-chain V192 O-N4V hydrogen bond, and a main-chain M194 NN3V hydrogen bond to the TDP’s 4V-aminopyrimidine ring. Conserved interactions linking the cofactor to the protein at the diphosphate end include ligation of an octahedrally coordinated Mg2+ ion by the side-chain oxygens of D230 and N260, the main-chain oxygen of Q262, a water oxygen, and two of the diphosphate oxygens. Other apparent protein–cofactor interactions, though not strictly conserved, include ring stacking between F602 and the 4Vaminopyrimidine ring, the hydrophobic side chain of M194 wedged underneath both cofactor rings, and H142 hydrogen bonds to the diphosphate moiety.

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Other nearby residues that do not interact directly with the cofactor but line the active-site cavity include H106, S109, Q140, Y177, E235, L264, and K392 from one subunit and D521, E522, I569, Y599, E636, and H640 from the ‘‘other’’ subunit. The active-site cavity has a wide mouth and deep cleft with dimensions of roughly 18 8 21 A˚. Based on their orientations and proximity to the location where cofactor–substrate adducts are expected to form, the residues E522, H640, Y177, and H106 are most likely to interact with reaction intermediates at some point(s) in the mechanism. As explained in the next section, H407, though unobserved in the crystal structure due to disorder, was later included in the list of active-site residues likely to be important in the reaction. Results from site-directed mutagenesis at these positions are presented later. A stereo picture illustrating the active site and cofactor binding is given in Figure 1c. C. Structural Comparison with Related Enzymes From a sequence homology point of view, E. coli PDHc-E1 is most similar (17% identity) to transketolase (TK), an enzyme operating on a diﬀerent substrate and having a very diﬀerent function in the pentose shunt. Both, however, are TDP dependent. A least-squares comparison of PDHc-E1 (8) and TK (9) crystal structures showed the two enzymes to be structurally very similar as well, with an rms deviation of 2.0 A˚ between 541 corresponding Ca atoms (10). The cofactor binding modes were also found to be similar, as was the conformation of the TDP. Based on the general structural similarity and the fact that the regions missing in E1 due to disorder were actually observed in TK, tentative positions for these regions in E1 at some point(s) in the reaction were deduced from a least-squares structural superposition. This superposition indicated that H407 in E1 (H263 in TK) could enter the E1 active site and be in a position to interact with substrate or reaction intermediates. Biochemical studies with substitutions at position 407 then conﬁrmed the importance of the histidine residue (10) and also suggested a role

Figure 1 (a) Ribbon drawings of the N-terminal, middle, and C-terminal domains (left to right) in the E. coli PDHc 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). (See color insert.)

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for it (to be discussed later) in both the TK and E1 reactions. The position of H407 in the active site as deduced from the least-squares superposition is shown in Figure 1c. A crystal structure for the TDP-dependent E1 component of a branched-chain dehydrogenase multienzyme complex (BDHc) with speciﬁcity for 2-oxoisovalerate has been reported (11), and it has also been compared to the E. coli PDHc-E1 structure (8). BDHc-E1 represents a diﬀerent class of E1 components, which form a2h2 heterotetramers rather than a2 homodimers. This structure has essentially no sequence homology with that of E. coli PDHc-E1, although parts of the reaction catalyzed are common and it serves a similar role in the overall multienzyme complex. Least squares structural alignment with PDHc-E1 was carried out, as in the comparison with TK, and resulted in an rms deviation of 2.12 A˚ between 404 corresponding Ca atoms. Despite the reasonable overall rms deviation, PDHc-E1 agreement with BDHc-E1 is signiﬁcantly worse than that with TK because substantially fewer matching atoms could be found. In addition, there are several places where major structural diﬀerences occur. Even when the least-squares alignment is carried out on individual domains independently, only about 50% of the Ca atoms could be matched. The unmatched regions correspond to large structural diﬀerences and are generally on the enzyme surface, suggesting possible diﬀerences in binding to other enzymatic components within the respective multienzyme complexes. Results from the least-squares alignments with TK and BDHc-E1 are shown in Figure 2a and 2b, respectively. III. MECHANISTIC INFORMATION ON THE E. coli PDHc-E1 A. Histidine Residues in the Active Center On one side of the TDP thiazolium ring, in proximity to the thiazolium C2 atom, a cluster of histidine residues, H106, H142, and H640, are found (Fig. 1c). Residue H142 is involved in TDP binding, forming a hydrogen bond with one of the oxygen atoms of the diphosphate moiety. Of these three histidines, residue H640 is located closest to the 4V-amino group of the 4V-aminopyrimidine ring. Residues H106 and H142 are located in the cleft between the two subunits leading to the TDP binding site. According to the PDHc-E1 crystal structure these histidine residues may be involved in pyruvate binding

Figure 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). (See color insert.)

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and/or stabilization of the covalent intermediate formed between pyruvate and TDP (8). To test this hypothesis, H106, H142, and H640 were converted to alanine by site-directed mutagenesis to create the corresponding E1 variants with single histidine substitutions. As shown in Table 1, the substitution of any of these three active-center histidine residues by itself to alanine led to only modest decreases in activity: (1) measured in the overall PDHc reaction after reconstitution with E2-E3 subcomplex (the activities were 10–26% as compared with parental E1); (2) measured with an E1speciﬁc activity assay using DCPIP to trap the enamine formed in the forward reaction with pyruvate (the activities were 32–75% as compared with parental E1); or (3) using HETDP as a substrate (the activities were 12–38% as compared with parental E1) (not presented). The single H106A, H142A, and H640A substitutions did not aﬀect signiﬁcantly the values of Km and Kd for TDP binding or the values of Km for pyruvate binding, indicating that these residues are not essential for the TDP and pyruvate binding (Table 1). The group at Rutgers has developed a MALDI-TOF mass spectrometric method to detect reductive acetylation of either a lipoyl domain expressed independently or the single-lipoyl domain from an E2 subunit (12) according to equations (2) and (3). The method not only obviates the need for radioactive pyruvic acid used earlier for the same purpose, but also oﬀers a rapid and precise method to identify substitutions at either E1 or E2 that impair the reductive acetylation (see later data in Sec. III. C). Under the conditions of the experiment, the lipoyl domain expressed independently was fully acetylated after 30 s of incubation with the PDHc-E1 and pyruvate (Figure 3A–C). The mass of the acetylated lipoyl domain was 9019 F 2 Da according to MALDI-TOF MS, compared with the theoretical mass of 9019 Da (lipoyl domain mass is 8975 Da, and it was increased by 44 mass units after the acetylation reaction). Reaction of the H106A, H142A, and H640A E1 variants with pyruvate and the lipoyl domain resulted in a mass of 9026 F 2.22 Da, indicating that the reductive acetylation of lipoyl domain is also unaﬀected by substitution at H106, H142, or H640. The foregoing results show that residues H106, H142, and H640 are not crucial for the decarboxylation step. The similar changes induced by the substitutions in the overall activity of the complex and in the model DCPIP reaction starting with HETDP (not presented) suggest that these histidine residues may be involved in the stabilization or subsequent reaction of the enamine intermediate. B. Residues Y177 and H179 Create a ‘‘ ‘‘Gate’’ ’’ That Regulates Intermediate Formation Extensive kinetic and spectroscopic data obtained in combination with the Xray structural data on the location of Y177 and H179 strongly suggest that

Information Transfer in the E. Coli PDHc Table 1

Activities and Kinetic Parameters for PDHc-E1 and Its Active-Center Variants

Substitution E1 H106A H142A H640A H179A Y177A Y177F H407A D521A E522A E571A E571D E571Q Y599A Y599F F602Y a

415

Pyruvate:NAD+ oxidoreductase activity (u/mg E1)a 18.0 (100%) 1.82 (10%) 4.75 (26%) 2.11 (12%) 1.72 (9.6%)c 0.99 (5.5%) 2.12 (12%)d 0.025 (0.14%) 0.80 (4.4%) 3.67 (20%) 0.19 (1%)e 0.32 (1.6%)e 0.68 (3.7%) 0.53 (3%) 8.80 (49%) 10.20 (57%)

Km pyruvate (mM )

DCPIP reaction, pyruvate (u/mg E1)a

Kd , TDP (lM )b

1.58

0.515

2.11

6.05

4.71

0.822

15.71

7.07

0.451

7.16

6.79

0.304

0.381 (100%) 0.284 (75%) 0.121 (32%) 0.191 (50%) 0.248 (65%) 0.087 (23%) 0.216 (57%) 0.047 (12%) 0.029 (7.6%) 0.137 (36%) 0.062 (16%) 0.132 (34%) 0.020 (5%) 0.052 (14%) 0.551 (145%) 0.242 (64%)

kcat (s1) 60.0

— 3.31

Km TDP (lM )

904.0 6.65

0.299 0.280

0.76

135.0

0.531

0.08

N/A

N/A

2.68

N/A

N/A

12.23

17.37

0.93

—

13.75

0.17

—

Not saturated 8.26

1.65

2.25 1.78

0.35

9.91

1.80

29.3

11.95

0.44

34.22

17.94

0.14

1.09 4.71 11.66 3.26 2.01 3.87 1.83 19.59 N/A 14.68 Not saturated 3.19 4.22 7.25 Not saturated

One unit of activity (u) is deﬁned as the amount of NADH produced (or DCPIP reduced in the E1speciﬁc reaction. b Dissociation constants (Kd) were determined by ﬂuorescence quenching. c The activity of the H179A E1 variant was measured in the presence of 0.2 mM TDP and may be enhanced at least sixfold once the TDP concentration is elevated to 5 mM. d The activity of the Y177F E1 variant was measured once the steady state was reached, since the progress curve for NADH production exhibited a prolonged lag phase. e The activity of the E571A and E571D E1 variants was measured in the presence of 0.20 mM TDP and may be enhanced two-fold (E571A E1) and 12-fold (E571D) once the TDP concentration is elevated to 5 mM.

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Figure 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. (See color insert.)

Y177 and H179 have a role in stabilization of the transition state(s) on the reaction pathway. According to the PDHc-E1 crystal structure, residue Y177 is located in the active-site cavity, with the tyrosine phenolic oxygen atom located at 6 A˚ from the thiazolium C2 atom, too far to assist in the ﬁrst essential deprotonation step required to trigger the reaction (Fig. 4). However, there is a water molecule located close to residue Y177 on the E1 subunit. This water molecule forms hydrogen bonds with the imidazole nitrogen atom of H640 and the N4V atom of the 4V-aminopyrimidine ring, and it is also near (3 A˚) the C2 atom of the thiazolium ring and to residue H106 (Figs. 1c and 4). This water molecule would almost certainly be replaced upon substrate binding. The Y177A and Y177F substitutions aﬀected the interaction of PDHcE1 with the E2-E3 subcomplex (the activities were 5.5% for Y177A, 12% for

Figure 3 (A) MALDI-TOF mass spectrum of the lipoyl domain. The spectrum shows the molecular ions for singly (mass = 8979 Da) and doubly (mass = 9183 Da) lipoylated forms of the lipoyl domain. (B) MALDI-TOF mass spectrum of acetylated lipoyl domain. The spectrum shows the acetylation of singly (mass = 9017 Da) and doubly (mass = 9222 Da) lipoylated forms of the lipoyl domain at 30 s of incubation with E1 (0.1 lM ) and pyruvate (2.0 mM ) in 20 mM KH2PO4 buﬀer (pH 7.0), containing 2 mM MgCl2, 0.2 mM TDP, and 0.6 mM lipoyl domain in a total volume of 0.1 mL at 25jC. (C) MALDI-TOF mass spectrum of the acetylated lipoyl domain at 30-min incubation with E1 and pyruvate.

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Y177F, and 9.6% for the H179A variant) and, to a lesser extent, the decarboxylation of pyruvate in the model reaction with DCPIP (the activities were 23% for Y177A, 57% for Y177F, and 65% for H179A) (Table 1). The progress curve for NADH production in the overall PDHc assay for both the Y177F and H179A variants revealed a prolonged lag phase, whose duration diminished upon increasing the TDP concentrations (Fig. 4 presents data for the H179A variant). The Km values for TDP were 135 lM (Y177F) and 904.0 lM (H179A), 85-fold and 570-fold higher than that of E1 parental (1.58 lM ), respectively (Table 1). However, according to the quenching of intrinsic E1 ﬂuorescence concomitant with TDP binding, both variants gave Kd values similar to that for parental E1 (Table 1). Also, the Y177A, Y177F, and H179A variants gave Km values for pyruvate similar to that for parental E1 at 0.20 mM TDP, indicating that substitution at Y177 and H179 does not aﬀect pyruvate binding (Table 1). The Y177 substitution in E1 did aﬀect the binding of thiamine 2-thiazolone diphosphate (TTDP, a TDP analog with a C2jO substitution at the thiazolium ring). According to reaction progress curve analysis, the Y177A substitution drastically changes the mechanism of inhibition of the Y177A E1 variant with TTDP as compared with the parental E1 (13). The Y177A substitution, however, does not aﬀect the binding of the thiamine 2-thiothiazolone diphosphate (TTTDP, a TDP analog with C2jS substitution at the thiazolium ring) according to ﬂuorescence and circular dichroism spectroscopic measurements (13). An important new result from these studies is the observation and characterization of a positive circular dichroism band in the 330-nm region upon addition of TTTDP (but not of TDP or TTDP) to parental E1 and the Y177A E1 variant (Fig. 5). We suggest that the band is the result of the formation of chiral TTTDP at the active center, chiral by virtue of the V-conformation enforced on all TDP enzymes studied to date (13), while TTTDP, but not TDP or TTDP, does have a weak UV absorption centered at 319 nm. The similar kinetic behavior of the H179A and Y177F variants of E1 located on the same loop in the E1 structure suggests that these residues and the loop on which both are located create a ‘‘gate’’ that regulates, perhaps locking in the intermediate, protecting it from side reactions. C. Histidine 407—A Residue Participating in the Reductive Acetylation of Both an Independently Expressed Lipoyl Domain and the Intact 1-lip E1 Subunit On the basis of the structural alignment described earlier, a disordered and unobserved region in PDHc E1 (residues 401–413) is likely to be structurally similar to the corresponding TK loop (residues 257–269) located near the

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Figure 5 Circular dichroism spectra of E. coli PDHc-E1 titrated with TTTDP. PDHc-E1 (9.0 lM ) in 10 mM KH2PO4 ( pH 7.0) was titrated with TTTDP at concentrations varying from 0.49 to 50 lM in the presence of 1 mM MgCl2. Inset: Dependence of the CD signal at 330 nm on the concentration of TTTDP.

TDP-binding region, with the H407 residue in PDHc-E1 corresponding to residue H263 in TK (10). A comparison of the E. coli PDHc-E1 and TK active centers also suggested that the potentially reactive H407 in E1 may indeed enter the E1 active site, as shown in Figure 1c, and participate in catalysis at some point(s) in the sequence of reactions. Accordingly, the H407A variant of E1 was created to probe whether H407 plays a role in the reaction mechanism. The activity of the H407A E1 variant measured in the overall PDHc reaction was approximately 0.14% relative to the parental E1 (Table 1). According to the E1-speciﬁc DCPIP assay, the speciﬁc activity of H407A E1 was about 12% starting with pyruvate (Table 1) and 17% starting with HETDP as substrates (data not presented), compared to the parental E1, indicating that the pathway through decarboxylations is only modestly aﬀected by the H407A substitutions. The value of Kd for TDP binding obtained from ﬂuorescence quenching was 1.83 lM, similar to that of the

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parental E1 (Table 1). In addition, there was the positive band at 330 nm produced in the near-UV CD spectrum of the H407A E1 variant on addition of TTTDP, indicating that binding of neither TDP nor TTTDP is aﬀected by the H407A substitution. The reduction in speciﬁc activity with the H407A substitution is similar to that reported for TK with the H263A substitution (14), indicating that both residues are involved in catalysis. To conﬁrm a hypothesis concerning the role of H407 in the interaction of PDHc-E1 with E2, the reductive acetylation of the lipoyl domain expressed independently and of the 1-lip E2 subunit by the parental E1 and its H407A variant were compared using MALDI-TOF mass spectrometry. The time course for reductive acetylation was diﬀerent for parental E1 and the H407A variant (Fig. 6); the lipoyl domain was fully reductively acetylated by parental E1 and pyruvate within 30 s (the ﬁrst time point), displaying a mass of 9019 F 2 Da for acetylated lipoyl domain. For the H407A E1 variant both forms of the lipoyl domain, unacetyalated (8975 Da) and acetylated, were observed during the time course (Fig. 6), indicating a slow rate of reductive acetylation. The calculated pseudo-ﬁrst-order rate constant was 0.0038 s1, signiﬁcantly smaller than the kcat of 0.80 s1 reported for E. coli E1 (15) and Azotobacter vinelandii E1 (16). Incubation of the H407A E1 variant and pyruvate with 1lip E2 displayed the presence of only the unacetylated form of lipoyl domain in the mass spectrum, indicating that H407 is important for reductive acetylation of the 1-lip E2 subunit. In addition, isothermal titration calorimetric measurements showed that binding of the lipoyl domain to parental E1 is exergonic (DGj = 6.6 kcal/mol), with a favorable enthalpy of binding (DHj = 0.62 kcal/mol) and an entropy of binding (T D S) of 5.98 kcal/mol, indicating that the interaction of the lipoyl domain with E1 is entropy driven. The Kd for binding lipoyl domain to E1 was 15 lM. A similar experiment carried out with the H407A E1 variant and the lipoyl domain revealed only very weak binding (DHj = 0.06 kcal/mol), indicating that H407 substitution in E1 also aﬀects the free energy of binding to the lipoyl domain. The parallel studies using lipoyl domain independently expressed and 1lip-E2 subunit clearly warn us that the former is just an appropriate chemical model for the intact E2 subunit and that the most signiﬁcant results are those obtained with the intact E2 subunit. On the basis of model studies carried out at Rutgers, which indicated that the lipoic acid needs to be activated by an electrophile to undergo a facile reaction (17,18), and the evidence presented earlier, we suggest that activation of the lipoamide covalently bonded to the E2 subunit toward reductive acetylation involves residue H407 of the E1 subunit (Scheme 2). According to Scheme 2, both TK and E1 + E2 carry out ‘‘ligation’’ reactions. More importantly from the point of view of catalysis, both must solve the diﬃcult

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Figure 6 Time course of reductive acetylation of the lipoyl domain by pyruvate and the H407A E1 variant monitored by MALDI-TOF mass spectrometry. Spectra show the unacetylated and acetylated forms of the lipoyl domain at 30 s, 3 min, 5 min, and 30 min of incubation of the H407A E1 variant (0.5 lM ) with pyruvate (2.0 mM ) in 20 mM HEPES (pH 7.0) containing 2 mM MgCl2, 0.2 mM TDP, and 0.6 mM of lipoyl domain in a total volume of 0.1 mL at 25jC.

problem of electrophilic activation of a very weak base, as seen in Scheme 2. In TK this takes the form of protonation of the carbonyl functional group, a ketone or aldehyde, while on E1-E2 it is the protonation of the dithiolane ring. The loop with H263 in TK is then predicted to have importance in the ligation step (14,19), just as H407 in E1 is important in the reductive acetylation step. Finally, we propose that the conserved features here seen will be

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Scheme 2

common to the entire TDP superfamily of enzymes charged with such ‘‘ligation’’ mechanisms. D. Experimental Evidence for Formation of the 4V-Iminopyrimidine Tautomeric Form of the Phosphonolactyl Thiamine Diphosphate in the Active Sites of the E. coli PDHc-E1 The compound phosphonolactylthiamine diphosphate (PLTDP) is the covalent adduct formed between methylacetylphosphonate (MAP) and TDP; it resembles LTDP, the ﬁrst covalent intermediate formed between the substrate and TDP on the PDHc reaction pathway (Scheme 1). It is a potential inhibitor of the E. coli PDHc-E1 (20) and binds reversibly in the active sites of PDHcE1; however, it will not undergo C–P bond cleavage, the reaction analogous to the decarboxylation step expected for LTDP (see Scheme 1). Fluorescence titration of PDHc-E1 with PLTDP led to a value of Kd = 2.69 lM, similar to the value of Kd = 2.11 lM for TDP binding. However, the near-UV CD spectrum exhibited signiﬁcant novel features in the 300–350-nm region on addition of PLTDP to the PDHc-E1, not easily seen with addition

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of TDP or HETDP (Fig. 7). The diﬀerence spectra of PDHc-E1 in the presence and absence of PLTDP revealed a positive CD peak centered at 305 nm (Fig. 7). The amplitude of this peak increased on addition of PLTDP and displayed saturation, providing a Kd = 0.71 lM (Fig. 7B inset), stronger than that determined by quenching of the intrinsic ﬂuorescence (Kd = 2.69 lM ). The CD feature here identiﬁed can be associated with the 1V,4ViminoTDP in the V-conformation at the active site of E1, chiral by virtue of the ﬁxed conformation. There is evidence from the Rutgers group that the positive circular dichroism peak observed at 305 nm most likely pertains to the 4V-iminopyrimidine tautomer of the PLTDP (21): In model compounds consisting of N1-methyl-4-aminopyrimidinium or N1-methyl-N4-n-butylpyrimidinium salts, on treatment with either NaOH in water, or 1,8-diazabicyclo[5.4.0]-undec-7-ene) as the base in dimethylsulfoxide, there is a new species formed with kmax in the ultraviolet spectrum near 307 and 302 nm, and this intermediate reverts back to the starting compound on acidiﬁcation. The 1H NMR chemical shifts are also consistent with the formation of the 1-methyl4-imino tautomer. On the E477Q active-center YPDC variant, the imino-TDP intermediate could be observed by rapid-scan stopped ﬂow with UV detection when mixing the enzyme with pyruvate in the presence of pyruvamide. Circular dichroism studies of the E477Q active-center YPDC variant in the presence of benzoylformic acid and pyruvamide revealed a positive CD signal at 306 nm that could be assigned to the imino-TDP tautomer as well and, importantly, ruling out the enamine as a possible source of this absorbance (since benzoylformic acid on decarboxylation should give rise to an enamine with Emax near 380 nm). Equipped with a number of PDHc-E1 active-site variants (activities and kinetic parametes of the E. coli active-site variants presented in Table 1), we could then interrogate the enzymes as to which amino acid is required for stabilizing the 1V,4V-imino TDP. Such studies are summarized in Table 2: 1. With three variants studied, F602Y, H640A, and Y599A, the band near 305 nm is in evidence. For F602Y E1 the CD signal displayed saturation with increasing PLTDP concentrations, similar to parental E1 (Kd = 0.76 lM ). However, for the H640A and Y599A variants of E1, no saturation was evident. Of these residues, F602 is stacked onto the 4V-aminopyrimidine ring of TDP, H640 is close enough to participate in hydrogen bonding to the substrate

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Figure 7 (A) Titration of PDHc-E1 with PLTDP (1–50 lM ). (B) Diﬀerence spectra resulting from subtraction of the spectrum of PDHc-E1. Inset: Variation of circular dichroism signal at 305 nm with PLTDP concentration.

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Table 2 Dissociation Constants and Circular Dichroism Maximum for PLTDP Binding to PDHc-E1 and Its Active-Center Variants

Substitution E1 F602Y H407A H640A Y599A E571A E571D a

Pyruvate:NAD+ oxidoreductase activity (%)

Kd, PLTDP ﬂuorescence (lM )

Circular dichroism maximum (nm)

Kd, PLTDP circular dichroism (lM )

100 57 0.14 12 3 1 1.8

2.69 5.33 9.96 4.06 3.83 2.13 19.5

305–306 308 — 304–305 306 — 305a

0.71 0.76 — Not saturated Not saturated — Not saturated

Maximum at 305 nm observed at concentrations of PLTDP higher than 35 lM.

and/or TDP-bound intermediates, and Y599 is in the channel leading to the active center. 2. With the H407A substitution, the CD band at 305 nm could not be detected. While no electron density was detected corresponding to this residue in the crystal structure of the E1, we have nevertheless shown that it has an important role in the reductive acetylation of E2 by E1 and pyruvate (see earlier). 3. Perhaps most importantly, the signal was not seen with the E571A substitution; residue E571 is within strong hydrogen-bonding distance of the N1V atom of TDP. This result strongly suggests that E571 has a role in stabilizing the 1V,4V-imino TDP tautomer. With the E571D variant, the negative charge is still present but is removed by approximately 1 A˚ from the N1V atom as compared to the parental enzyme. Accordingly, the CD signal at 305 nm is still seen, albeit at a much higher concentration of PLTDP, and the signal does not display saturation binding (Table 2). Also, neither the E571D nor the E571A variant displays saturation with TDP (Table 1). In summary, residues E571 and H407 appear to have the most dramatic eﬀects on the appearance of the CD band assigned to the 1V,4V-iminoPLTDP, while the other three substitutions do inﬂuence the binding according to the inability to be saturated by this compound. We conclude from the model studies, the PDHc-E1 with PLTDP results, and those recently reported with the YPDC E477Q variant, that the presence of the 1V,4V-iminoTDP on TDP enzyme is now established to the best of our

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ability. Whether or not it can be detected by a particular experimental method depends on the variant and the TDP analog. Equally signiﬁcantly, so far as PLTDP is a suitable analog for LTDP at the active center, we conclude that in the ﬁrst substrate–TDP covalent tetrahedral adduct, there is a preponderance of the 1V,4V-imino TDP tautomer, perhaps a result of steric crowding, almost certainly poised to participate in catalysis. Finally, we suggest that the Breslow mechanism for TDP catalysis be augmented with inclusion of a role(s) for the 4V-aminopyrimidine and its 1V,4V-imino tautomer. These ﬁndings make TuDP an astonishingly versatile coenzyme, unique in providing both electrophilic (via the thiazolium ring) and acid-base (via the 4V-amino/4Vimino tautomeric equilibration) catalytic activity. E. Partitioning and Utilization of the LTDP and HETDP Intermediates in the PDHc-E1 Reaction According to Scheme 1 and the mechanism of the PDHc-E1 reaction, LTDP is the ﬁrst intermediate formed between the C2 atom of TDP and pyruvate. LTDP undergoes decarboxylation in the active sites of the PDHc-E1 forming 2-a-hydroxyethylidene-TDP, the enamine intermediate common to all TDPdependent enzymes. In the PDHc reaction, the enamine intermediate would be oxidized by the lipoic acid covalently amidated onto the E2 component. As already mentioned, an external artiﬁcial oxidizing agent, DCPIP, can be used to trap the enamine formed, either in the forward reaction, starting with pyruvate (to demonstrate possible impairment of the reaction through decarboxylation), or starting with HETDP, testing whether the C2a–H bond can be ionized for the subsequent oxidation/reduction process. The E. coli PDHc or its E1 component can be reconstituted with LTDP and shows 0.03% of activity in the overall PDHc reaction (kcat = 0.0196 s1), compared to pyruvate as substrate (kcat = 60.0 s1), and 9% in the DCPIP model reaction (kcat = 0.115 s1), compared to pyruvate as substrate (1.27 s1). The low activity observed in the presence of LTDP in both reactions may be explained by the slow release of the TDP formed in the active sites of E1 with LTDP as a substrate. In addition, it was shown that E1 catalyzed the decomposition of LTDP in the reverse reaction, leading to formation of pyruvate and TDP, as detected with a lactate dehydrogenase/NADH coupled assay (kcat for the reverse reaction was 0.078 s1, as compared with the kcat = 0.115 s1 obtained in the forward reaction and detected by DCPIP). HETDP is formed by protonation of the enamine at the C2-a position, a reaction most likely not on the PDHc pathway. Yet it has been used as substrate in the overall PDHc reaction and in the DCPIP model reaction (Table 3), although the activities are much lower than those with the natural substrates (pyruvate-TDP). The activity, compared to the natural substrates,

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Table 3 Comparison of Pyruvate-TDP and HETDP as Substrates for PDHc-E1 According to the Overall PDHc Assay and DCPIP Reaction Pyruvate:NAD+ oxidoreductase, pyruvate-TDP (s1) 60.0 (100%)

Pyruvate:NAD+ oxidoreductase, HETDP (s1)

2,6-DCPIP, pyruvate-TDP (s1)

2,6-DCPIP, HETDP (s1)

2,6-DCPIP, HETDPa,h-d4 (s1)

0.040 (0.066%)

1.27 (100%)

0.150 (11.8%)

0.072

diminished more in the overall reaction (0.066%) than in DCPIP reaction (11.8%). This is likely due to the fact that the binding of HETDP and the release of TDP is more diﬃcult in the overall reaction with PDHc. It was also shown by MALDI-TOF mass spectrometry that the lipoyl domain expressed independently (at 0.6 mM concentration) is acetylated by HETDP (0.2 mM) and parental E1, with the rate much smaller than with pyruvate as substrate (kapp = 0.0025 s1 as compared to 0.8 s1). Comparison of the rates with E1 and DCPIP, HETDP, and HETDP-d4 displayed a small primary kinetic isotope eﬀect for kcat (Dkcat = 2.08), which indicates that proton abstraction is only partially rate limiting (Table 3). In conclusion, when using HETDP as substrate, we are presenting evidence for generation of the enamine according to: The DCPIP trapping reaction, and more importantly, the observation of NADH production derived solely from HETDP. This is a remarkable reaction, since HETDP is not believed to be on the reaction pathway in the reductive acetylation of the lipoyl-E2 by hydroxyethylidene-E1. It demonstrates yet once more the ability of TDP-dependent decarboxylases to stabilize zwitterionic intermediates, already demonstrated for yeast pyruvate decarboxylase in the laboratory at Rutgers (22,23). The kinetic isotope eﬀect, while modest, suggests that C2-aH ionization of HETDP is partially rate limiting, again aﬃrming that enamine formation from HETDP proceeds reductive acetyl transfer. F. Some Evidence Regarding the Negative Circular Dichroism Band Near 320 nm The near-UV circular dichroism spectra of the TDP-dependent enzymes yeast TK and pigeon pyruvate dehydrogenase (E1) are widely used to study the interaction of these enzymes with TDP (14,24,25). The binding of the TDP to the yeast TK is accompanied by the appearance of a negative circular dichroism band with a maximum at 320 nm, a diagnostic for TDP binding (14,24). From our experience with the E. coli PDHc-E1, addition of diﬀerent concentrations of TDP to the PDHc-E1 is not accompanied by any signiﬁ-

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cant changes in the region of 300–350 nm of the CD spectrum (13). In the presence of 0.2 mM pyruvate to assist the TDP binding to PDHc-E1 and at saturating ThDP concentration (200 lM ), weak positive changes were observed in the 300 to 330-nm region of the CD spectra, similar to those observed for PLTDP binding, more than likely indicative of the formation of the 1V,4V-imino TDP (Fig. 8). However, the diﬀerence spectra obtained after subtraction of the spectrum of PDHc-E1 in the absence of TDP does not show the nicely shaped peak at 305–306 nm detected with PLTDP binding. Subsequent addition of 2 mM pyruvate to PDHc-E1 saturated with TDP led to the immediate appearance of a stable negative band with a maximum at 320–330 nm, similar to that observed for the yeast TK (26) and E477Q YPDC in recently published data from the Rutgers laboratory (21). This band at 320– 330 nm was also observed with the F602Y E1 and D521A E1 variants. It is almost certainly the case that some form of TDP, either ionization state or tautomeric state, is responsible for this CD signal, observed now on three diﬀerent TDP enzymes (two sources of E1, TK, and YPDC). In the case of PDHc-E1 and YPDC, we suggest that the observation reﬂects that state of

Figure 8 Titration of PDHc-E1 with TDP and pyruvate. 1: baseline; 2: spectrum of PDHc-E1 (20 lM ) in the absence of TDP; 3: spectrum of PDHc-E1 in the presence of 0.20 mM TDP, 2 mM MgCl2, and 0.2 mM pyruvate; 4: conditions as in spectrum 3, with 2 mM pyruvate added.

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the enzymes in which a ‘‘ligation’’ type of reaction takes place. With the PDHc-E1, only addition of pyruvate produced the signal, while with YPDC the signal was seen with the E477Q variant, which behaved as an acetoin synthase.

IV. SUMMARY AND CONCLUSIONS The crystal structure of the PDHc-E1 from E. coli has been determined at 1.85-A˚ resolution. This represents the ﬁrst structural example of an E1 component from the a2 homodimeric class and also the ﬁrst example of a functional E1 of any sort with speciﬁcity for pyruvate. The structure has enabled identiﬁcation of residues in the active site likely aﬀecting catalysis, and comparisons with a related a2h2 heterotetrameric BDHc-E1 as well as with TK are made. The latter comparison enabled identiﬁcation of an additional conserved residue, H407, also important in catalysis. The mechanism of the reaction taking place exclusively on E1 and then the reductive acetylation of lipoyl domain per se and of the single lipoyl domain E2 have been examined. 1. Substitution of any of the four histidines H106, H142, H179, and H640 appears to have no dramatic eﬀect on the E1-speciﬁc reactions, while H640 may have a modest eﬀect on reductive acetylation. 2. Substitution of Y177 appears to aﬀect intermediate formation, and the similar kinetic behavior of H179 and Y177 on the same loop suggest that these residues create a ‘‘gate’’ that regulates intermediate formation. 3. Reconstitution of PDHc-E1 with LTDP and HETDP demonstrated that these intermediates are converted to the expected products by E1, enabling assignment of speciﬁc roles to the active-center residues. Especially the utilization of HETDP in reductive acetylation strongly supports the Rutgers hypothesis that a major role for TDP enzymes is to stabilize zwitterionic intermediates. 4. The most informative substitution to date is at H407. The H407A variant has activity nearly equal to that of parental enzyme in E1speciﬁc assays; but there is clear impairment of the reductive acetylation from a variety of experiments. 5. Several of the E1 variants gave clear evidence for the formation of a TDP conformation that gives rise to the often observed negative CD band centered near 330 nm. It has become evident that at least some of the residues too mobile to be seen in the crystal structure of the PDHc-E1 subunit have important functions

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in catalysis, such as the residue histidine 407. Further studies on the interaction of the PDHc-E1 and PDHc-E2 subunits promise to provide important insight in to both intersubunit information transfer and assembly of the entire complex.

ACKNOWLEDGMENTS It is a pleasure to acknowledge a very fruitful collaboration with Professor John Guest of Sheﬃeld University, UK, who initially provided the molecular biological constructs to help us initiate this program. Supported at Pittsburgh by NIH-GM-61791 (to WF) and at Rutgers by NIH-GM-62330 and the NSF Training Grant -BIR 94/13198 in Cellular and Molecular Biodynamics (F.J.).

ABBREVIATIONS TDP TTTDP TTDP HETDP LTDP PLTDP MAP CD DCPIP MALDI-TOF MS PDHc E. coli PDHc-E1 E2

E3 YPDC TK BDHc-E1

thiamine diphosphate thiamine 2-thiothiazolone diphosphate thiamine 2-thiazolone diphosphate C2-a-hydroxyethylthiamine diphosphate 2-a-lactylthiamine diphosphate phosphonolactylthiamine diphosphate methylacetylphosphonate circular dichroism 2, 6-dichlorophenolindophenol matrix-assisted laser desorption ionization time-of-flight mass spectrometry pyruvate dehydrogenase multienzyme complex Escherichia coli pyruvate dehydrogenase, the first subunit of PDHc (TDP dependent) dihydrolipoamide acetyltransferase with a single lipoyl domain per subunit, the second subunit of PDHc dihydrolipoamide dehydrogenase, the third subunit of PDHc yeast pyruvate decarboxylase transketolase branched-chain dehydrogenase multienzyme complex E1 subunit

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REFERENCES 1.

2. 3. 4. 5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

M Koike, LJ Reed, WR Carroll. a-Keto acid dehydrogenation complexes. I. Puriﬁcation and properties of pyruvate and a-ketoglutarate dehydrogenation complexes of Escherichia coli. J Biol Chem 235:1924–1930, 1960. IC Gunsalus. The Mechanism of Enzyme Action. Johns Hopkins University Press, Baltimore, MD, 1954, pp. 545–580. V Massey. The Enzymes 2nd ed. Academic Press, New York, 1963, pp. 275–305. LJ Reed. Multienzyme complexes. Acc Chem Res 7:40–46, 1974. PE Stephens, MG Darlison, HM Lewis, JR Guest. The pyruvate dehydrogenase complex of Escherichia coli K12. Nucleotide sequence encoding the pyruvate dehydrogenase component. Eur J Biochem 133:155–162, 1983. PE Stephens, MG Darlison, HM Lewis, JR Guest. The pyruvate dehydrogenase complex of Escherichia coli K12. Nucleotide sequence encoding the dihydrolipoamide acetyltransferase component. Eur J Biochem 133:481–489, 1983. PE Stephens, HM Lewis, MG Darlison, JR Guest. Nucleotide sequence of the lipoamide dehydrogenase gene of Escherichia coli K12. Eur J Biochem 135:519– 527, 1983. 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. Y Lindqvist, G Schneider, V Ermler, M Sundstrom. Three-dimensional structure of transketolase, a thiamin diphosphate–dependent enzyme at 2.5-A˚ resolution. EMBO J 11:2373–2379, 1992. N Nemeria, P Arjunan, A Brunskill, F Sheibani, W Wei, Y Yan, S Zhang, F Jordan, W Furey. Histidine 407, a phantom residue in the E1 subunit of the Escherichia coli pyruvate dehydrogenase complex, activates reductive acetylation of lipoamide on the E2 subunit. An explanation for conservation of active sites between the E1 subunit and transketolase. Biochemistry 41:15459–15467, 2002. A Aevarsson, K Seger, S Turley, J Sokatch, W Hol. Crystal structure of 2oxoisovalerate dehydrogenase and the architecture of 2-oxo acid dehydrogenase multienzyme complexes. Nature Struct Biol 6:785–792, 1999. W Wei, H Li, N Nemeria, F Jordan. Expression and puriﬁcation of the dihydrolipoamide acetyltransferase and dihydrolipoamide dehydrogenase subunits of the Escherichia coli pyruvate dehydrogenase multienzyme complex: a mass spectrometric assay for reductive acetylation of dihydrolipoamide acetyltransferase. Protein Expression Puriﬁcation 28:140–150, 2003. N Nemeria, Y Yan, Z Zhang, A Brown, P Arjunan, W Furey, JR Guest, F Jordan. Inhibition of the Escherichia coli pyruvate dehydrogenase complex E1 subunit and its tyrosine 177 variants by thiamin 2-thiazolone and thiamin 2thiothiazolone diphosphate. J Biol Chem 49:45969–45978, 2001. C Wikner, U Nilsson, L Meshalkina, C Udekwu, Y Lindqvist, G Schneider. Identiﬁcation of catalytically important residues in yeast transketolase. Biochemistry 36:15643–15649, 1997. D Graham, LC Packman, RN Perham. Kinetic and speciﬁcity of reductive

432

16.

17.

18.

19.

20.

21.

22.

23. 24.

25.

26.

27. 28.

Furey et al. acetylation of lipoyl domain from 2-oxo acid dehydrogenase multienzyme complexes. Biochemistry 28:1574–1581, 1989. A Berg, AH Westphal, A De Kok. Kinetic and speciﬁcity of reductive acetylation of wild-type and mutated lipoyl domains of 2-oxo-acid dehydrogenase complexes from Azotobacter vinelandii. Eur J Biochem 252:45–50, 1998. CC Chiu, A Chung, G Barletta, F Jordan. Intramolecular model for the reductive acyl transfer catalyzed by the a-keto acid dehydrogenase. J Am Chem Soc 118: 11026–11029, 1996. 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. 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. R Kluger, DC Pike. Active-site-generated analogues of reactive intermediates in enzymic reactions. Potent inhibition of pyruvate dehydrogenase by a phosphonate analogue of pyruvate. J Am Chem Soc 99:4504–4506, 1977. 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. Bioorganic Chemistry 30:188–198, 2002. F Jordan, H Li, A Brown. Remarkable stabilization of zwitterionic intermediates may account for a billion-fold rate acceleration by thiamin diphosphate– dependent decarboxylases. Biochemistry 38:6369–6373, 1999. M Liu, Z Zhang, F Jordan. To be published. L Meshalkina, C Wikner, U Nilsson, Y Lindqvist, G Schneider. Biochemistry and Physiology of Thiamin Diphosphate Enzymes. A.u.C. Intemann, Wissenschaftlicher Verlag, Prien, Germany, 1996, pp. 532–542. LG Korochkina, LS Khailova, SE Severin. Investigation of the pyruvate dehydrogenase complex using the circular dichroism method. Biokhimiya 56: 1840–1849, 1991. E Fiedler, R Golbik, G Schneider, K Tittmann, H Neef, S Ko¨nig, G Hu¨bner. Examination of donor substrate conversion in yeast transketolase. J Biol Chem 276:16051–16058, 2001. P Kraulis. MOLSCRIPT: A program to produce both detailed and schematic plots of protein structures. J Appl Crystallogr 24:946–950, 1991. M Carson. RIBBONS 2.0. J Appl Crystallogr 24:958–961, 1991.

24 Structure, Function, and Regulation of Pyruvate Dehydrogenase Kinase Kirill M. Popov and Alina Tuganova University of Missouri–Kansas City, Kansas City, Missouri, U.S.A. Mellissa M. Bowker-Kinley, Boli Hung, Pengfei Wu, C. Nicklaus Steussy, Jean Hamilton, and Robert A. Harris Indiana University School of Medicine, Indianapolis, Indiana, U.S.A.

I. INTRODUCTION The pyruvate dehydrogenase complex functions as the enzymatic link between glycolysis and the citric acid cycle. Using NAD+ and CoA as cosubstrates and thiamine pyrophosphate, lipoic acid, and FAD as prosthetic groups, the pyruvate dehydrogenase complex carries out oxidative decarboxylation of pyruvate to give acetyl-CoA, NADH, and CO2. Although ﬂux through the pyruvate dehydrogenase complex is critically important for complete disposal of glucose, almost complete shutdown of the complex is necessary in the fasting state to conserve the three carbon compounds (lactate, pyruvate, and alanine) required for glucose synthesis. This is because the thiamine pyrophosphate–dependent decarboxylation of pyruvate by the complex is irreversible and glucose cannot be synthesized from two-carbon compounds. A mechanism must therefore be turned on during fasting that induces nearly complete inhibition of the pyruvate dehydrogenase complex 433

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in most cells of the body. Although valuable in starvation, this same mechanism is unfortunately also activated in diabetes and thereby contributes to the hyperglycemia. The central role in inactivation of the pyruvate dehydrogenase complex belongs to pyruvate dehydrogenase kinase, which catalyzes the sequential phosphorylation of three serine residues of the a-chain of the E1 component, rendering the entire complex inactive. In humans, four genetically and biochemically diﬀerent forms of pyruvate dehydrogenase kinase, designated as PDK1, PDK2, PDK3, and PDK4 (1–3), are found. Over the years approximately 25 genes of a PDK lineage have been identiﬁed in various eukaryotic species. Analysis of these genes shows that each protein molecule consists of two parts almost equal in length: a poorly conserved amino-terminal half, and a highly conserved carboxy-terminal half. The topology of one of the members, PDK2, is shown in Figure 1. Within the carboxyl terminus there are at least four regions, which are uniformly conserved among all these proteins: boxes N, G1, G2, and G3. The N box is deﬁned by an invariant Asn residue. The G1 box is deﬁned by the consensus sequence -Asp-X-Gly-X-Gly-. The G2 and G3 boxes are deﬁned by the consensus sequences -Gly-X-Gly-X-GlyLeu- and -Gly-X-Gly-Thr-. This arrangement is characteristic of PDK-

Figure 1 Topology of rat PDK2. The cylinders represent a-helices and the arrows h-strands. The helices are numbered a1 through a11 and the sheets h1 through h7. The conserved boxes, identified from sequence alignment with the histidine kinases, are also shown.

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related proteins, strongly suggesting that this class of protein kinases is not related to the Ser/Thr- and Tyr-protein kinases residing in other cellular compartments. In mitochondria, the enzymatic activity of pyruvate dehydrogenase kinase is regulated by an array of metabolites. In general, the products of the dehydrogenase reaction (NADH and acetyl-CoA) activate, while the substrates (pyruvate, NAD+, and CoA) inhibit kinase activity (4). This allows the kinase molecule to sense the metabolic need for carbohydrate catabolism and to adjust the activity of the pyruvate dehydrogenase complex according to existing demands. This rather short-term regulation adjusts ﬂux through the pyruvate dehydrogenase complex on a minute-to-minute basis. In addition there is long-term regulation. This term refers to the hyperphosphorylated state of pyruvate dehydrogenase that develops under conditions such as starvation, diabetes, ischemic heart disease, sepsis, lactic acidosis, and high-fat feeding. In contrast to the short-term eﬀects, long-term eﬀects correlate with the stable increase in pyruvate dehydrogenase kinase activity. In this chapter we will discuss recent developments from our laboratories on (1) the analysis of the structure and function of the pyruvate dehydrogenase kinase and (2) the long-term regulation of pyruvate dehydrogenase kinase expression. II. THREE-DIMENSIONAL STRUCTURE OF PYRUVATE DEHYDROGENASE KINASE The three-dimensional structure of pyruvate dehydrogenase kinase (isozyme PDK2) complexed with ADP was recently solved at 2.8-A˚ resolution (5). The structure revealed a homodimeric arrangement (Fig. 2). The monomer of PDK2 consists of two well-deﬁned subdomains corresponding to the amino and carboxyl termini of the kinase molecule. The amino-terminal domain consists of six a-helices and one h-strand connected by ﬂexible loops (Figs. 1 and 2). Overall the amino-terminal domain is dominated by four amphipatic a-helices arranged as a four-helix bundle. The hydrophobic surfaces of each helix form the interface that stabilizes the structure of the bundle. Hydrophilic surfaces are exposed to the solvent. The amino terminus winds around the four-helix bundle, forming multiple contacts with side chains of amino acids from the four-helix bundle, thus stabilizing the overall structure of the aminoterminal domain. The carboxy-terminal domain is folded as a mixed a/h sandwich (Figs. 1 and 2). The sandwich consists of six h-strands and four a-helices. The ﬁrst layer of the sandwich is assembled from ﬁve h-strands organized in a slightly twisted h-sheet. The second layer of the sandwich is built of four a-helices, which are positioned almost parallel to the corresponding h-strands. A small

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antiparallel h-sheet seals the sandwich at one end. A hydrophobic core stabilizes the overall structure of the domain. The carboxy-terminal and amino-terminal domains (Fig. 2C and D) are connected by a ﬂexible, poorly ordered loop. They are stabilized relative to each other by a small antiparallel h-sheet, as well as through multiple hydrophobic interactions. The two monomers in the dimer are in a ‘‘head-to-head’’ orientation. The dimer interface is formed by the h-sheets of the carboxyterminal domain. The interface buries 2200 A˚2 of the molecule and is stabilized by a number of hydrophobic interactions and hydrogen bonds between the monomers. The remainder of the molecule extends out into a horseshoe shape (Fig. 2B), with the two amino-terminal domains as the arms and the carboxy-terminal interface at the base. III. MECHANISM OF NUCLEOTIDE BINDING Electron density corresponding to the bound ADP was found at the center of the cone-shaped cavity of the carboxy-terminal domain. The cavity is topped by a loop reminiscent of a ‘‘lid,’’ which likely controls access of ATP into the pocket (6). The lid appears to be disordered, indicating mobility of this region. The distal part of the lid curves around the a- and h-phosphate groups of the ADP, forming a structure reminiscent of the P-loop of other nucleotidebinding proteins (6). Multiple hydrophobic side chains projecting up from the face of the h-sheet form the base of the pocket. As a result, the adenine ring is buried against a hydrophobic surface created by these residues. The binding site is unusual in that it provides few direct polar contacts with the nucleotide and instead makes a substantial number of solvent-mediated interactions. Some of these interactions involve water molecules with low-crystallographictemperature factors. Interaction between the adenine base and the protein (Fig. 3A) involves a direct hydrogen bond from the C6 amino group (N6) of adenine to the carboxylate side chain of Asp282 (G1 box). In addition, Asp282 provides one of the three protein ligands for the water molecule that donates a hydrogen bond to the N1 imino nitrogen of the adenine. Two other protein ligands for this

Figure 2 Overall architecture of the PDK2 complexed with magnesium-ADP. Two subunits of the PDK2 dimer (shown as a ribbon diagram), each containing separate kinase and regulatory domains. The dimer associates along the h-sheets of the kinase domain. (A) View of the dimer with the noncrystallographic twofold axis in the plane of the paper. (B) PDK2 dimer viewed down the noncrystallographic twofold axis. (C) Ribbon representation of the carboxy-terminal kinase domain. (D) Ribbon representation of the amino-terminal regulatory domains.

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Figure 3 Interactions in the nucleotide–binding pocket of PDK2. Schematic diagram showing the interactions of the adenine ring and ribose moiety (A) and of the polyphosphate tail (B). Residues from the protein and ADP are drawn as stick representations. Water molecules are shown as spheres and hydrogen bonds as broken rods.

water molecule come from the side chain of the invariant Thr354 (G3 box) and the peptide nitrogen of the invariant Gly286 (G1 box). These interactions might be critical in determining the strict speciﬁcity of PDK2 for adenyl nucleotides. A second hydrogen bond to adenine N6 is provided by another water molecule, which is bound by the peptide carbonyl of Leu244. The N7 of the adenine is bound to a third water molecule, attached through the extensive bonding network formed by the water molecules and the backbone of the polypeptide. The 2V-hydroxyl of the ribose makes a direct hydrogen bond to the side chain of an invariant Thr302 of the ATP-lid. The carbonyl oxygen of Thr302

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is bound to the 3V-hydroxyl of the ribose, which in turn forms a hydrogen bond with the side chain of the invariant Arg250 that protrudes across the nucleotide-binding pocket. The position of Arg250 is stabilized through bonding with Glu254. The phosphate moiety of ADP is bound to PDK2 by numerous direct interactions as well as by interactions mediated by a well-ordered Mg2+ ion (Fig. 3B). The side chain of Asn247 (N box) donates a hydrogen bond to the a-phosphate group. On the opposite side, the a- and h-phosphate groups are surrounded by the amino acids of the B-loop (Gly317, Phe318, Gly319, Tyr320, Gly321, and Leu322). Nitrogen atoms of Phe318, Gly319, and Leu322 on the P-loop donate a hydrogen bond to an oxygen atom of a phosphate. This allows the phosphate groups to tuck in close to the end of the helix a4 (Figs. 1 and 3B). An octahedrally coordinated Mg2+ ion links all phosphates to the protein. Each phosphate provides an oxygen atom to chelate the Mg2+ ion. Analysis of the three-dimensional structure clearly shows that the invariant amino acids (boxes N, G1, G2, and G3) identiﬁed through sequence alignment are strategically positioned to participate in nucleotide binding (Fig. 4). The side chain of the invariant Asn247 (N box) binds directly to the h-phosphate of ATP and to the magnesium ion. Asp282 (G1 box) interacts with the N6 atom in the adenine base. Gly286 (G1 box) allows the main chain

Figure 4 Proposed mechanism of the phosphotransfer reaction, involving nucleophilic attack by the hydroxyl group of Ser264 on the g-phosphate group of ATP with Glu243 acting as general base.

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to contour the adenine ring and serves as a protein ligand for the water molecule bound to the N1 imino nitrogen of the adenine. Gly319 (G2 box) is part of the P-loop that allows the main chain to make a sharp turn in the vicinity of the h- and g-phosphate groups and donates a hydrogen bond to an oxygen atom of a h-phosphate. Finally, the invariant Thr354 (G3 box, consensus -G351-x-Gly353-Thr354-Asp355-) serves as a third protein ligand for the water molecule, donating a hydrogen bond to the N1 imino nitrogen of the adenine. The functional signiﬁcance of Asn247, Asp282, Gly286, and Gly319 has been analyzed by site-directed mutagenesis (7). Four mutant kinases, Asn247A, Asp282A, and Gly286A lacked any appreciable activity, whereas Gly319A was found to be catalytically active but diplayed approximately 100 times greater Km value for ATP. ATP-binding analysis showed that Asn247A, Asp282A, and Gly286A could not bind nucleotide. The Gly319A mutant bound ATP so poorly that it was diﬃcult to determine the actual binding constant. The results of this functional analysis are completely consistent with available structural information. IV. CATALYTIC MECHANISM OF PYRUVATE DEHYDROGENASE KINASE Although pyruvate dehydrogenase kinase does not resemble Ser/Thr- and Tyr-speciﬁc protein kinases, its protein fold is not novel. It is remarkably similar to the structure of histidine kinases that are widely distributed in bacteria. The archetypal histidine kinase is a dimer displaying a characteristic modular structure that consists of a central dimerization domain folded as a four-helix bundle and two nucleotide-binding domains arranged around the dimerization domain (8). Although somewhat diﬀerent in connectivity, both the dimerization and nucleotide-binding domains of the histidine kinases ﬁt well with the amino- and carboxy-terminal domains of pyruvate dehydrogenase kinase. Histidine kinases catalyze the ATP-dependent phosphorylation of a conserved histidine residue using a trans-autophosphorylation mechanism whereby one monomer phosphorylates the other monomer within the dimer. The phospho-accepting histidine residues are provided by the dimerization domain and serve as intermediates in the phospho-transfer pathway, accepting phosphoryl groups from ATP and transferring them to downstream targets. To serve as phospho-acceptor, the conserved histidine residues are positioned on the outside surfaces of the four-helix bundle and are solvent exposed. Quite remarkably, pyruvate dehydrogenase kinase possesses an invariant histidine residue (His115 or H box in PDK2) readily identiﬁable through sequence alignment with the histidine kinases. Based on this observation it was suggested that pyruvate dehydrogenase kinase might use circuitry similar to that of the histidine kinases to phosphorylate pyruvate dehydrogenase.

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In agreement with this idea, earlier studies have established that treatment of pyruvate dehydrogenase kinase with histidine-modifying reagents leads to a marked inhibition of kinase activity (9). However, when His115 was altered to Ala in rat PDK2 using site-directed mutagenesis, the resulting mutant kinase was fully active (M. M. Bowker-Kinley and K. M. Popov, unpublished ﬁnding). This indicates that there are structural factors preventing His115 from serving as acceptor of a phospho group. Analysis of the three-dimensional structure of pyruvate dehydrogenase kinase shows that the helix carrying the invariant histidine residue is ‘‘kinked’’ at Pro120. This causes His115 to adopt a conformation in which the imidazole ring is hidden inside the four-helix bundle, where it forms a bond to the side chain of Ser83. Thus structural studies, along with the results of site-directed mutagenesis, strongly suggest that it is highly unlikely that pyruvate dehydrogenase kinase uses a phosphohistidine intermediate to catalyze phospho-transfer. The lack of support for the existence of a phospho-enzyme intermediate in the reaction catalyzed by pyruvate dehydrogenase kinase prompted us to explore the hypothesis that pyruvate dehydrogenase kinase uses a general base catalytic mechanism to activate the hydroxyl group of the serine residue of the protein substrate for the direct attack on g-phosphate group (10). The nucleotide-binding domain of PDK2 contains the invariant Glu243, which is ideally located to act as a general base and promote nucleophilic attack. Furthermore, His239, located on the next turn of the helix, is positioned to polarize Glu243. The crystal structure of PDK2 also contains a water molecule hydrogen-bonded to Glu243. This water molecule might occupy the position of the hydroxyl group of the serine residue when the nucleotide-binding site contains bound ADP. In order to test this idea, we used site-directed mutagenesis to convert Glu-243 to Ala, Asp, or Gln and His-239 to Ala. The resulting mutant kinase demonstrated a greatly reduced capacity for phosphorylation of pyruvate dehydrogenase. The Glu-243-to-Asp mutant had approximately 2% residual activity, while the Glu-243-to-Ala or -GLN mutants exhibited less than 0.5% or 0.1% residual activity, respectively. Activity of the His-239-to-Ala mutant was decreased by about 90%. Active-site titration with [a-32P]ATP revealed that neither the Glu-243 nor His-239 mutations aﬀected nucleotide binding. All mutant kinases showed similar aﬃnity to the wild-type kinase for the pyruvate dehydrogenase complex. Furthermore, pyruvate dehydrogenase kinase was found to possess a weak ATP hydrolytic activity, which required Glu-243 and His-239, similar to the kinase activity. These observations strongly suggest that Glu243 might act as a general base, which activates the hydroxyl group for direct attack on the g-phosphate group. His-239 has a role in aligning and polarizing the glutamate residue. Thus, it appears that acatalytic domain of this type can support transfer of a g-phosphate to either serine or histidine residues. The latter is possible

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because of minor but fundamental diﬀerences in the amino acid composition of the catalytic domains of pyruvate dehydrogenase kinase and the histidine kinases. In histidine kinases, phosphorylation occurs as a result of direct attack on the g-phosphate by a histidine residue, which is part of the dimerization domain. The nucleophilicity of the attacking histidine residue is likely to increase through interaction with neighboring residues. Accordingly, histidine kinases lack the amino acids equivalent to Glu-243 and His-239 of pyruvate dehydrogenase kinase. In striking contrast, pyruvate dehydrogenase kinase cannot utilize His-115 for phospho-transfer because it is hidden within a four-helix bundle. Instead, it uses a general base catalysis mechanism to activate the serine residue of a protein substrate for direct attack on the g-phosphate group of ATP. V. REGULATION OF THE PYRUVATE DEHYDROGENASE COMPLEX ACTIVITY BY ALTERED EXPRESSION OF ITS KINASE Much of the pyruvate dehydrogenase complex is inactive in tissues of animals that have been starved or made diabetic. The inhibition of the complex is the result of the activity of the pyruvate dehydrogenase kinases phosphorylating multiple serines in the E1 alpha subunit. Inactivation of the complex by this means is physiologically important because it helps conserve substrates that can be converted to glucose, which the brain continuously needs for the synthesis of ATP. This mechanism sets in when blood levels of free fatty acids increase during starvation and diabetes. Greater availability promotes fatty acid oxidation in almost every tissue of the body except the brain. This invariably increases the mitochondrial concentrations of NADH and acetylCoA, which indirectly stimulate pyruvate dehydrogenase kinase activity by reduction and acetylation of the lipoyl groups of E2. Although it will require more time to establish this, a second mechanism is possible that results in an even greater increase in pyruvate dehydrogenase kinase activity. This mechanism involves a stable increase in pyruvate dehydrogenase kinase activity that is independent of the activating eﬀects of acetyl-CoA and NADH on the kinase (11,12). Until recently it was not clear whether this mechanism involved covalent modiﬁcation of the kinase or an increase in its amount. The ﬁnding that four pyruvate dehydrogenase kinase isozymes are expressed in tissues (1–3) motivated studies to examine whether altered expression of one or more of these isozymes might be responsible. This turned out to be the case. Starvation of rats for food and induction of diabetes by chemical destruction of the pancreatic h-cells have been found to increase the level of PDK4 expression in heart (Fig. 5), skeletal muscle, liver, kidney, and lactating mammary gland (13–17). These conditions also cause a more modest increase

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Figure 5 The relative amount of PDK isoenzymes 1, 2, and 4 in heart mitochondria of well-fed, starved, and streptozotocin-diabetic rats. Rats (200 g of body weight, n = 4) were either well fed, 48-hour fasted, or treated with 65 mg/kg streptozotocin prior to sacrifice. PDK isoenzymes were quantified in mitochondrial extracts by Western blot analysis with antisera specific for PDK1, 2, and 4. Known amounts of the corresponding recombinant isoenzymes were loaded to standardize the signals provided by the antisera.

in the level of PDK2 expression in several of these tissues. Refeeding and insulin therapy completely reverse the eﬀects of starvation and diabetes on PDK4 and PDK2 expression in these tissues, respectively (13–17). High dietary fat (18,19), carnitine deﬁciency (20), and hibernation (21) also increase PDK4 expression in tissues. Hypoxia down-regulates PDK4 expression in the heart (22), the only tissue thus far examined, presumably to promote glucose and pyruvate oxidation and therefore to maximize the eﬃciency of oxygen utilization by the heart in this condition (22). The hormonal and nutritional factors involved in regulation of PDK4 and PDK2 expression are now of great interest and are beginning to be deﬁned. Insulin plays an important role, as evidenced by its eﬀectiveness in decreasing expression of both PDK4 and PDK2 in diabetic animals. Glucocorticoids are candidate regulatory molecules, since the concentrations of these compounds increase signiﬁcantly in starvation and diabetes. Indeed, treatment of rats with the synthetic glucocorticoid dexamethasone causes a rapid increase in PDK4 expression in a number of tissues (M. M. BowkerKinley, P. Wu, and R. A. Harris, unpublished ﬁndings). This suggests that

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corticosterone is important in the rat and cortisol in the human for induction of PDK4 expression during starvation and diabetes. Likewise, injection of rats with WY-14,643, a synthetic mimic of fatty acids and also a relatively speciﬁc PPARa agonist, causes a rapid increase in PDK4 expression in many tissues (14). WY-14,643 loses its eﬀectiveness in PPARa-null mice, indicating that WY-14,643 exerts its eﬀects on PDK4 expression via this receptor. The increase in PDK4 expression that normally occurs in response to starvation is attenuated in PPARa-null mice, suggesting that activation of this receptor is involved in PDK4 expression in response to starvation. Based on the known activation of PPARa by free fatty acids and the increase in blood levels of free fatty acids that occurs in response to starvation, diabetes, carnitine deﬁciency, hibernation, and high-fat feeding, we have hypothesized that activation of PPARa by free fatty acids may be involved in promoting PDK4 expression (23,24). The importance of glucocorticoids, insulin, and free fatty acids has been further investigated in studies carried out with Morris hepatoma 7800C1 cell and L6 myotubes. The cells of these model systems are rich in mitochondria, and these cells retain sensitivity to stimulation by glucocorticoids and insulin. Indeed, incubation of Morris hepatoma 7800C1 cells (23) and L6 muscle cells (M. M. Bowker-Kinley and R. A. Harris, unpublished) with dexamethasone causes a rapid increase in PDK4 expression as measured by changes in PDK4 message and protein. No eﬀect on message stability is observed, suggesting dexamethasone stimulates transcription of the PDK4 gene. The eﬀect occurs at physiologically important concentrations of glucocorticoids and is inhibited by RU486, indicating involvement of the glucocorticoid receptor. Less than an hour is required for a signiﬁcant increase in PDK4 message level. Thus, the increase in glucocorticoids in starvation and diabetes is surely one of the factors involved in increasing PDK4 expression in major tissues of the body. It is therefore likely that the hyperglycemia characteristic of Cushing syndrome can be explained in part by the increase in PDK4 expression caused by the elevated blood levels of glucocorticoids in this disease. Insulin very eﬀectively blocks and reverses induction of PDK4 expression by dexamethasone in both 7800C1 hepatoma cells and L6 cells. The ability of insulin to dominate over glucocorticoids in this regard is similar to the negative eﬀect that insulin has on glucocorticoid induction of phosphoenolpyruvate carboxykinase in hepatoma cells (25). PDK2 is constitutively expressed in 7800C1 hepatoma cells and L6 cells. Glucocorticoids are without eﬀect upon PDK2 expression in these cells. However, insulin very eﬀectively down-regulates PDK2 expression in both cell types. The negative eﬀects of insulin on PDK4 and PDK2 expression are time and concentration dependent; however, PDK4 levels decrease much faster in response to insulin

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because the half-lives of the PDK4 message and PDK4 protein are very short. The eﬀects of insulin on PDK4 and PDK2 expression are mediated at the levels of transcription and message stability, respectively (B. Huang and R. A. Harris, unpublished studies). Since WY-14,643 is a relatively speciﬁc ligand for PPARa, we have proposed activation of this nuclear receptor/transcription factor promotes PDK4 expression. Since free fatty acids activate PPARa (24), it follows that the increase in free fatty acids occurring in starvation and diabetes may stimulate PDK4 expression via PPARa in these conditions. Consistent with this idea, both WY-14,643 and free fatty acids are eﬀective inducers of PDK4 expression in 7800C1 hepatoma cells (23). In contrast, WY-14,643 alone causes very little increase in PDK4 expression in L6 myotubes (M. M. Bowker-Kinley and R. A. Harris, unpublished ﬁndings). However, the combination of dexamethasone and WY-14,643 induces a much greater increase in PDK4 expression than either compound alone, an apparent synergism likely explained by up-regulation of PPARa by the glucocorticoids (26). Since it is likely that WY-14,643 induces its eﬀects by activation of PPARa and since this receptor can also be activated by free fatty acids (27), the eﬀects of free fatty acids on PDK4 expression have been examined in 7800C1 hepatoma cells. Palmitate and oleate are both eﬀective at physiologically relevant concentrations in inducing PDK4 expression as measured at the message and protein levels. These ﬁndings suggest that activation of PPARa by free fatty acids is involved in promoting PDK4 expression in starvation and diabetes. ACKNOWLEDGMENTS This work is supported by U.S. Public Health Service Grants DK 47844 (RAH) and GM 51262 (KMP), American Diabetes Association Grant (JAH and CNS), American Heart Association Grant-in-Aid (PW), the Grace M. Showalter Residuary Trust, predoctoral fellowships from the Indiana University School of Medicine Center for Diabetes Research (BH), and the American Heart Association (MMB-K). ABBREVIATIONS PDC PDK E1 E2 PPARa

pyruvate dehydrogenase complex pyruvate dehydrogenase kinase pyruvate dehydrogenase dihydrolipoamide transacetylase peroxisome proliferator-activated receptor a

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REFERENCES 1.

2.

3.

4.

5.

6. 7. 8. 9.

10.

11.

12.

13.

14.

15.

16.

KM Popov, NY Kedishvili, Y Zhao, Y Shimomura, DW Crabb. Primary structure of pyruvate dehydrogenase kinase establishes a new family of eukaryotic protein kinases. J Biol Chem 268:26602–26606, 1993. R Gudi, MM Bowker-Kinely, NY Kedishvili, Y Zhao, KM Popov. Diversity of the pyruvate dehydrogenase kinase gene family in humans. J Biol Chem 270: 28989–28994, 1995. MM Bowker-Kinley, WI Davis, P Wu, RA Harris. Evidence for existence of tissue-speciﬁc regulation of the mammalian pyruvate dehydrogenase complex. Biochem J 329:191–196, 1998. S Ravindran, GA Radke, JR Guest, TE Roche. Lipoyl domain–based mechanism for the integrated feedback control of the pyruvate dehydrogenase complex by enhancement of pyruvate dehydrogenase kinase activity. J Biol Chem 271:653–662, 1996. CN Steussey, KM Popov, MM Bowker-Kinley, RB Jr, Sloan, RA Harris, JA Hamilton. Structure of pyruvate dehydrogenase kinase. Novel folding pattern for a serine protein kinase. J Biol Chem 276:37443–37450, 2001. R Dutta, M Inouye. GHKL, an emergent ATPase/kinase superfamily. Trends Biochem Sci 25:24–28, 2000. MM Bowker-Kinley, KM Popov. Evidence that pyruvate dehydrogenase kinase belongs to the ATPase/kinase superfamily. Biochem J 344:47–53, 1999. AM Bilwes, LA Alex, BR Crane, MI Simon. Structure of CheA, a signaltransducing histidine kinase. Cell 96:131–141, 1999. BP Mooney, NR David, JJ Thelen, JA Miernyk, DD Randall. Histidinemodifying agents abolish pyruvate dehydrogenase kinase activity. Biochem Biophys Res Commun 267:500–503, 2000. A Tuganova, MD Yoder, KM Popov. An essential role of Glu-243 and His-239 in the phosphotransfer reaction catalyzed by pyruvate dehydrogenase kinase. J Biol Chem 276:17994–17999, 2001. GS Denyer, AL Kerby, PJ Randle. Kinse activator protein mediates longer-term eﬀects of starvation on activity of pyruvate dehydrogenase kinase in rat liver mitochondria. Biochem J 239:347–354, 1986. MC Sugden, MJ Holness, E Donald, H Lall. Substrate interactions in the shortand long-term regulation of renal glucose oxidation. Metabolism 48:707–715, 1999. P Wu, J Sato, Y Zhao, J Jaskiewicz, KM Popov, RA Harris. Starvation and diabetes increase the amount of pyruvate dehydrogenase kinase isoenzyme 4 in rat heart. Biochem J 329:197–201, 1998. P Wu, K Inskeep, MM Bowker-Kinley, KM Popov, RA Harris. Mechanism responsible for inactivation of skeletal muscle pyruvate dehydrogenase complex in starvation and diabetes. Diabetes 48:1593–1599, 1999. P Wu, PV Blair, J Sato, J Jaskiewicz, KM Popov, RA Harris. Starvation increases the amount of pyruvate dehydrogenase kinase in several mammalian tissues. Arch Biochem Biophys 381:1–7, 2000. MC Sugden, A Kraus, RA Harris, MJ Holness. Fibre-type speciﬁc modiﬁcation

Structure, Function, and Regulation of PDK

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

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of the activity and regulation of skeletal muscle pyruvate dehydrogenase kinase (PDK) by prolonged starvation and refeeding is associated with targeted regulation of PDK isoenzyme 4 expression. Biochem J 346:651–657, 2000. SJ Peters, RA Harris, GJF Heigenhauser, LL Spriet. Muscle ﬁber–type comparison of pyruvate dehydrogenase kinase activity and isoform expression in fed and fasted rats. Am J Physiol 280:R661–R668, 2001. SJ Peters, RA Harris, TL Pehleman, GJF Heigenhauser, LL Spriet. Human skeletal muscle PDH kinase activity and isoform expression during three days of a high-fat/low-carbohydrate diet. Am J Physiol. 281:E1151–E1158, 2001. MJ Holness, A Kraus, RA Harris, MC Sugden. Targeted up-regulation of pyruvate dehydrogenase kinase (PDK)-4 in slow-twitch skeletal muscle underlies the stable modiﬁcation of the regulatory characteristics of PDK induced by highfat feeding. Diabetes 49:775–781, 2000. M Horiuchi, K Kobayashi, M Masuda, H Terazono, T Saheki. Pyruvate dehydrogenase kinase 4 mRNA is increased in the hypertrophied ventricles of carnitine-deﬁcient juvenile visceral steatosis (JVS) mice. Biofactors 10:301–309, 1999. MT Andrews, TL Squire, CM Bowen, MB Rollins. Low-temperature carbon utilization is regulated by novel gene activity in the heart of a hibernating mammal. Proc Natl Acad Sci USA 95:8392–8397, 1998. P Razeghi, ME Young, S Abbasi, H Taegtmeyer. Hypoxia in vivo decreases peroxisome proliferator-activated receptor a-regulated gene expression in rat heart. Biochem Biophys Res Commun 287:5–10, 2001. B Huang, P Wu, MM Bowker-Kinley, RA Harris. Regulation of pyruvate dehydrogenase kinase expression by PPARa ligands, glucocorticoids, and insulin. Diabetes 51:276–283, 2002. P Wu, J Peters, RA Harris. Adaptive increase in pyruvate dehydrogenase kinase 4 during starvation is mediated by peroxisome proliferator-activated receptor a. Biochem Biophys Res Commun 287:391–396, 2001. K Sasaki, TP Cripe, SR Koch, TL andrenone, DD Petersen, EG Beale, DK Granner. Multihormonal regulation of phosphoenolpyruvate carboxykinase gene transcription. The dominant role of insulin. J Biol Chem 259:15242–15251, 1984. T Lemberger, B Staels, R Saladin, B Desvergne, J Auwerx, W Wahli. Regulation of the peroxisome proliferator-activated receptor a gene by gluococorticoids. J Biol Chem 269:24527–24530, 1994. H Keller, C Dreyer, J Medin, A Mahfoudi, K Ozato, W Wahi. Fatty acids and retinoids control lipid metabolism through activation of peroxisome proliferator-activated receptor-retinoid X receptor heterodimers. Proc Natl Acad Sci USA 90:2160–2164, 1993.

Figure 25.1 Crystal structure of the human E1b heterotetramer at 2.7-A˚ resolution. Each 400-residue a subunit (in purple and blue) is composed of a large domain containing mostly a-helices. The a subunit also shows an extended N-terminal tail and a small helical C-terminal domain. Each h subunit (in red and yellow) of 346 residues is divided in two domains of similar size. They correspond to the N-terminal and Cterminal halves of the polypeptide. Each domain of the h subunit comprises four parallel central h-strands, with several helices packed in variable orientations against the h-strands. The interactions between the ah and aVhV heterodimeric intermediates result in the assembly of the 167-kDa E1b heterotetramer. Each human E1b binds two TDP molecules (yellow spheres) at a–hV and aV–h subunit interfaces. (From Ref. 12.)

Figure 25.2 Cofactor TDP-binding sites in human E1b. Schematic representation of the cofactor binding site. Gln112-a and protein ligands of the magnesium ion have been omitted for clarity. (From Ref. 12.)

Figure 25.5 Mutations aﬀecting subunit interactions between a and hV subunits of human E1b. Mutations in the small C-terminal domain of the a subunit at the interface with the two h subunits. The C-terminal domain is shown with a coil representation of the backbone with selected side chains. The h subunits are shown as semitransparent surfaces, in yellow for the hV subunit in red for the h subunit. Some side chains in the h subunits that are forming the interface to the a subunit are shown beneath the surface. The a subunit makes contacts with both h subunits, with the prevalence of aromatic residues at the subunit interface. The sites of the three MSUD mutations in this region of the a subunit, F364C-a, Y368C-a, and Y393N-a, are highlighted. All three mutations are substitutions of aromatic residues involved in the interface to a h subunit. Tyr368-h is in contact with the h subunit (a–h interface), whereas Phe364-a and Tyr393-a contact the hV subunit (a–hV interface). (From Ref. 12.)

Figure 25.7 Nucleotide-binding pocket of rat BCK. Shown is superimposed 2Fo–Fc simulated annealing, with electron density maps (green, contoured at 1.5j) calculated for potassium (cyan) and magnesium (yellow) as well as bound ADP (A) and ATPgS (B). An anomalous diﬀerence Fourier electron density map (red, contoured at 5j) is shown for the ATPgS-bound BCK structure. The side chain of Phe303 has been removed for clarity. (From Ref. 13.)

Figure 25.8 Superposition of the apo-structure of rat BCK (red) with its ADP-bound (blue) ATPgS-bound forms (green). Nucleotide (ADP or ATPgS) binding orders H302 and F336 (electron density not visible in the apo-form) in the B/K domain interface to form a quadruple aromatic stack between Y301, H302, F336, and F338. Structural diﬀerences in the BH3 and BH4 helices between the apo and the nucleotide-bound states are depicted. (From Ref. 13.)

25 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 University of Texas Southwestern Medical Center, Dallas, Texas, U.S.A.

I. INTRODUCTION The mammalian mitochondrial branched-chain a-ketoacid dehydrogenase (BCKD) complex catalyzes the oxidative decarboxylation of branched-chain a-ketoacids derived from amino acids leucine, isoleucine, and valine (1). Genetic defects in this 4 106-dalton macromolecular complex result in the accumulation of branched-chain amino and a-ketoacid acids, leading to the heritable maple syrup urine disease (MSUD). The clinical phenotype of classic MSUD includes an early onset of often-fatal acidosis, with neurological derangement and mental retardation in survivors (2). The mammalian BCKD complex is organized around a cubic core of 24-meric dihydrolipoyl transacylase (E2b), to which multiple copies of branched-chain a-ketoacid decarboxylase (E1b), dihydrolipoamide dehydrogenase (E3), BCKD kinase (abbreviated as BCK), and BCKD phosphatase are attached through ionic interactions (3,4). The activity of the BCKD complex is tightly regulated by a 449

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phosphorylation/dephosphorylation cycle in response to dietary and hormonal signals (5). Phosphorylation of Ser-292 and Ser-302 in the a subunit of E1b by BCK results in the inactivation of the BCKD complex (6), whose activity is restored through dephosphorylation of the covalently modiﬁed E1b by BCKD phosphatase. The macromolecular architecture and reaction mechanisms for the BCKD complex are very similar to those for the related pyruvate and a-ketoglutarate dehydrogenase complexes. The three multienzyme systems make up the highly conserved family of mitochondrial a-ketoacid dehydrogenase complexes. X-ray and NMR structures for the components and domains of prokaryotic a-ketoacid dehydrogenase complexes have been elucidated (7– 11). In contrast, three-dimensional structures for components and domains of mammalian a-ketoacid dehydrogenase complexes are largely unknown. Our laboratory has recently determined crystal structures for thiamine diphosphate (TDP)–dependent E1b (12) and its regulatory enzyme BCK (13), as well as the solution structure for the single lipoyl-bearing domain (LBD) (14) of the mammalian BCKD complex. The structural information to be provided in this chapter constitutes the ﬁrst step toward understanding the organization and macromolecular interactions of the mammalian BCKD complex at the atomic level. Moreover, this knowledge provides a structural basis for the mechanism by which MSUD mutations disrupt structure/function and macromolecular assembly of the human BCKD complex.

II. CRYSTAL STRUCTURE OF HUMAN BRANCHED-CHAIN A-KETOACID DECARBOXYLASE (E1b) AND STRUCTURAL BASIS FOR MAPLE SYRUP URINE DISEASE (MSUD) A. Subunit Assembly and the TDP-Binding Pocket of E1b The E1b component catalyzes the TDP-mediated decarboxylation of branched-chain a-ketoacids and the subsequent reductive acylation of the oxidized lipoyl moiety covalently attached to E2b. The mammalian E1b component contains two a and two h subunits, with Mr = 45,500 and 37,500, respectively, forming an a2h2 heterotetramer. The E1 component has a size of Mr = 166,600 and requires K+ and Mg2+ ions and TDP for activity (15). The crystal structure of human E1b has recently been determined at 2.7-A˚ resolution (12). The overall a2h2 heterotetrameric structure of human E1b is shown in Figure 1. The tetrahedral arrangement dictates that each subunit be in contact with the other three subunits. Subunits a, aV, h, and hV are designated such that a and h subunits when combined correspond to one polypeptide of the related homodimeric yeast transketolase (16). Prominent features in the structure include the crossover of the N-terminal tails of the

Figure 1 Crystal structure of the human E1b heterotetramer at 2.7-A˚ resolution. Each 400-residue a subunit (in purple and blue) is composed of a large domain containing mostly a-helices. The a subunit also shows an extended N-terminal tail and a small helical C-terminal domain. Each h subunit (in red and yellow) of 346 residues is divided in two domains of similar size. They correspond to the N-terminal and Cterminal halves of the polypeptide. Each domain of the h subunit comprises four parallel central h-strands, with several helices packed in variable orientations against the h-strands. The interactions between the ah and aVhV heterodimeric intermediates result in the assembly of the 167-kDa E1b heterotetramer. Each human E1b binds two TDP molecules (yellow spheres) at a–hV and aV–h subunit interfaces. (From Ref. 12.) (See color insert.)

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two a subunits and the C-terminal extensions of these subunits that provide interactions with the h subunits. The small C-terminal extension of the a subunit is critical for the a2h2 assembly of human E1b and is the site of the prevalent Mennonite mutation (Y393N-a) (see later). The TDP cofactors are bound in the two active sites of the heterotetramer at the aV–h and a–hV interfaces (Fig. 2). The side chains of Gln112-a, Tyr113-a, Arg114-a, Arg220-a, and His291-a and the main-chain atoms of Gly194-a and Ala195-a in the a subunit are in direct contact with the oxygen atoms of the phosphate groups of the cofactor (Fig. 2). The bound Mg2+ ion is octahedrally coordinated between the cofactor phosphates, the carbonyl group of Tyr224-a, as well as the side chains of Asn222-a and Glu193-a. The side chain of Tyr102-hV is packed against one side of the aminopyrimidine ring of the cofactor, with the side chain of Leu164-a approaching the other side of the ring and wedged in between the two ring systems of the cofactor. Side chains of Ile226-a and Leu74-hV also contribute to cofactor binding through hydrophobic interactions. Glu76-hV is directly bound to the N1V atom, and the carbonyl group of Ser162-a is in contact with the N4V group on the opposite side of the aminopyrimidine ring. These two interactions, as well as the coordination of the Mg2+ ion and the conformation of the cofactor (torsion angles fr = 100j, fp = 71j in the human E1b structure), are the conserved features among proteins of known structure in the TDP superfamily. B. The Novel Structural K+ Ions The K+ ion bound at site 1 stabilizes a loop containing residues 161–166 of the a subunit (Fig. 3). Residues in this loop are well conserved among E1 proteins of a-ketoacid dehydrogenase complexes, including Ser161-a, Thr166-a, and Gln167-a, which interact directly with the metal ion through their side chains. Other atoms involved in the binding of the metal ion are main-chain carbonyl oxygens of Ser161-a and Pro163-a. The structural integrity of the K+ ion–binding loop 161–166 is apparently critical for the activity of the enzyme. The residues of this loop contribute directly to cofactor binding by hydrophobic interactions, which occurs through the above-mentioned Leu164-a situated between the two rings of the TDP, and by hydrogen bonding between the carbonyl group of Ser162-a and the N4V amino group of the cofactor (Fig. 3). Both of these interactions are conserved among TDPutilizing proteins with known structure (8,17–20). The binding of the K+ ion at this position to the h-chain of E1b explains the observations that enzyme activity as well as stabilization of the protein by TDP are dependent on the presence of K+ ions. The second K+ binding site is found in the h subunit at the interface with the small C-terminal domain of the a subunit (data not shown). The binding of a K+ ion at this site is important for the structural integrity of E1b at the a–h subunit interface.

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Figure 2 Cofactor TDP-binding sites in human E1b. Schematic representation of the cofactor binding site. Gln112-a and protein ligands of the magnesium ion have been omitted for clarity. (From Ref. 12.) (See color insert.)

C. The Structural Basis for MSUD There are currently over 25 known MSUD-causing mutations that lead to amino acid substitutions in the a or h subunit of human E1b (2). With a few exceptions, mutations in the E1b subunits often result in the manifestation of severe classic MSUD phenotype. Based on the human E1b structure, MSUDrelated missense mutations can be classiﬁed into three groups: The ﬁrst group

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Figure 3 Potassium site 1 (a subunit). The metal stabilizes a loop involved in cofactor binding. The metal ion is bound by two main-chain carbonyl groups and by the side chains of Ser161-a, Thr166-a, and GLn167-a. The side chain of Leu164-a and the main-chain carbonyl group of Ser162-a make direct contact with the TDP cofactor. (From Ref. 12.)

includes mutations that impede TDP-cofactor binding; the second group aﬀects mainly hydrophobic cores; and the third group occurs at the subunit interfaces, thereby interfering with interactions between a and h subunits. In the ﬁrst group are several MSUD mutations that aﬀect TDP-cofactor binding. Asn222-a is an invariant residue in the consensus sequence motif for TDP-dependent enzymes (21) and contributes to cofactor binding by coordinating to the bound Mg2+ ion (Fig. 4). The strict conservation of Asn222-a within the family of TDP-dependent enzymes is consistent with the critical function of this residue. This function apparently cannot be performed by a serine residue, because the N222S-a mutation causes the severe classic form of MSUD. It is plausible that in wild-type human E1b, the side-chain carbonyl oxygen of Asn222-a is used for binding the Mg2+ ion, whereas the hydroxyl group of the serine side chain is a poorer ligand to metal ions. Moreover, the Og of a serine residue is not able to extend as far toward the Mg2+ ion as the Oy1 of the asparagine. The combined eﬀects result in markedly reduced aﬃnity of the Asn222S-a mutant E1b for TDP, as reﬂected by a 103-fold

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Figure 4 MSUD mutations aﬀecting cofactor binding in human E1b. The R114W-a, R220W-a, and N222S-a mutations each aﬀect residues that are directly involved in binding the cofactor phosphates and the associated magnesium ion. Asn222-a is a conserved residue in the superfamily of TDP-utilizing enzymes and is a part of its sequence ﬁngerprint motif (GDG(X)22–30NN). Also shown is the primary phosphorylation site at Ser292-a. (From Ref. 12.)

increase in the KM value as compared with the wild-type enzyme (data not shown). The R114W-a and R220W-a classic MSUD mutations likewise have severe impacts on the cofactor binding. Both mutations introduce a tryptophan residue in the place of the arginine residue that coordinates to the phosphate groups of TDP through ionic interactions (Fig. 4). Substitutions with the bulky tryptophan residue at positions 114 and 220 not only abolish these interactions but also probably prevent cofactor binding by steric hindrance. Given the major eﬀects expected on TDP binding and possibly substrate binding due to these mutations, it is not surprising that these mutations result in the classic severe type of MSUD. A portion of the hydrophobic core in the a subunit is formed by side chains from helices 3 and 11 and strands a, d, e, h, and i. Two group 2 mutations in the a subunit presumably disrupt this hydrophobic core. The large charged side chain of arginine introduced by the T265R-a mutation in

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helix 11 cannot be accommodated within this core in the altered conformation. The much less severe I281T-a mutation located in strand i is very close to Thr265-a. The I281T-a substitution introduces a polar side chain, which may not be easily tolerated in this exclusively hydrophobic environment. In the family of heterotetrameric E1 proteins (about 50 known sequences so far), only leucine, methionine, isoleucine, and valine are found at this position. Patients heterozygous for I281T-a/Q145K-a manifest an intermediate clinical phenotype, with cells derived from these patients showing signiﬁcant residual enzyme activity. Two other mutations that also appear to prevent the formation of an optimal hydrophobic core are G204S-a and A208T-a. Both mutations introduce a larger polar side chain in an environment in which a smaller and nonpolar residue is required. The M64T-a mutation in helix 2 points to a critical location in a hydrophobic core in the a subunit, where four helices come together through hydrophobic interactions. Replacement of this methionine residue, which is invariant within the family of heterotetrameric E1b proteins, by the polar and smaller side chain of a threonine apparently has an adverse eﬀect on E1 stability. Residual multienzyme-complex activity in cells from patients carrying this mutation is also signiﬁcant, giving rise to an intermediate clinical phenotype. The phenotype is in agreement with the expected limited eﬀect of the methionine-to-threonine substitution in this position on E1b stability. There are three group 3 MSUD missense mutations that map to the extended small C-terminal domain protruding from the bulk of the a subunit (Fig. 5). This domain has diﬀerent sizes in various E1 heterotetramers and is in contact with both h and hV subunits in human E1b (Fig. 5). The interface contacts are to a large extent hydrophobic in nature, with a notable array of aromatic side chains from the a subunit side-packed against a host of aromatic residues from the hV subunit. All three MSUD mutations in this part of the a subunit, or the so-called ‘‘Mennonite region,’’ substitute an aromatic residue for a smaller and nonaromatic residue. The name for this region is based on the frequent tyrosine-to-asparagine mutation at position 393-a found in the U.S. Mennonite population (22,23), in which the prevalence of this mutation is 1 in 176 live births as a result of consanguinity (24,25). Tyr393-a is packed in between two aromatic residues, Trp330-hV and Phe324hV, of the hV subunit and forms two hydrogen bonds via its hydroxyl group with Asp328-hV as well as His385-a (Fig. 5). The tyrosine-to-asparagine mutation at position 393-a introduces a much smaller polar asparagine side chain, which is apparently unable to compensate for the extensive contact made by the native tyrosine side chain. The side chain of Tyr368-a, another residue in the ‘‘Mennonite region’’ aﬀected by MSUD mutations, ﬁts into a surface pocket on the h subunit, which is formed by the side chains of residues Asn183-h, Lys161-h, and Pro259-h and the main-chain atoms of Gly159-h and Ile160-h (Fig. 5). The hydroxyl group is able to act as both hydrogen-

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Figure 5 Mutations aﬀecting subunit interactions between a and hV subunits of human E1b. Mutations in the small C-terminal domain of the a subunit at the interface with the two h subunits. The C-terminal domain is shown with a coil representation of the backbone with selected side chains. The h subunits are shown as semitransparent surfaces, in yellow for the hV subunit and in red for the h subunit. Some side chains in the h subunits that are forming the interface to the a subunit are shown beneath the surface. The a subunit makes contacts with both h subunits, with the prevalence of aromatic residues at the subunit interface. The sites of the three MSUD mutations in this region of the a subunit, F364C-a, Y368C-a, and Y393N-a, are highlighted. All three mutations are substitutions of aromatic residues involved in the interface to a h subunit. Tyr368-h is in contact with the h subunit (a–h interface), whereas Phe364-a and Tyr393-a contact the hV subunit (a–hV interface). (From Ref. 12.) (See color insert.)

bond acceptor and donor by making hydrogen bonds to a main-chain carbonyl group (position 160-h) and a main-chain amino group (Gly159h). These speciﬁc interactions apparently cannot be formed by the smaller cysteine side chain, as a result of the tyrosine-to-cysteine mutation at position 368-a. Phe364-a, the third member of the aromatic array which is changed into a nonaromatic in certain MSUD patients, makes extensive hydrophobic

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interactions with the side chains of Leu363-a, Leu375-a, Gln37-a, and Leu382-a, which contribute to the compact hydrophobic core within the small C-terminal domain of the a subunit. In addition, these residues are also part of the interface to the h subunit, where the side chains of Leu362-a, Leu363-a, and Phe364-a are tightly packed against the side chain of Tyr313hV (Fig. 5). The phenylalanine-to-cysteine MSUD mutation at position 364-a is likely to aﬀect a–hV subunit interactions; it might also have profound consequences for the folding of the small C-terminal domain itself.

III. CRYSTAL STRUCTURE OF THE RAT BCKD KINASE A. Overall Structural Organization Crystal structures of rat BCK in its apo-form, in complex with ADP and with ATPgS have been determined at resolutions of 2.98 A˚, 2.29 A˚, and 2.7 A˚, respectively. These structures uniformly show a homodimeric arrangement, with each monomer comprising two domains (Fig. 6A). The ﬁrst domain (referred to as the B domain, residues 1–186) contains two parallel helix– hairpin–helix motifs that form a four-helix bundle (helices BH1, BH2, BH3, and BH4). The ﬁrst helix in each motif is interrupted by a kink. The second domain (referred to as the K domain, residues 187–373) contains a ﬁvestranded h-sheet (strands KB1, KB2, KB3, KB4, and KB5) opposed by a layer of three helices (KH1, KH2, and KH3) and harbors the nucleotidebinding site. A long loop (residues 303–339) that surrounds part of the nucleotide-binding site is disordered to varying degrees, depending on the bound nucleotide. The B and K domains are connected by a ﬂexible linker (residues 182–194), which adopts diﬀerent conformations in the BCK structures. The B and K domains within a BCK monomer interact with each other through two interfaces (Fig. 6A). The ﬁrst interface is near the linker region and is formed by residues close to the N-terminus (amino acids 39–42) or the C-terminus (amino acids 373–374). A second, more intimate interface is located between helices BH4 and KH3 in close proximity to the nucleotidebinding pocket. It contains a hydrophobic cluster and is formed through aromatic stacking interactions (Ala140, Leu160, Tyr301, His302, and Phe338), an ionic interaction between Arg157 and Asp300, as well as two polar interactions between the side chains of Asp161 and Tyr301 and between the carbonyl oxygen of Ala140 and the side chain of His302. The domain interface buries a total of about 2580 A˚2 of the surface area. The BCK dimer in our crystal structure consists of two crystallographically related molecules that interact through the concave surfaces of their K domains (Fig. 6B). This mode of dimer formation has never been described previously for any kinase containing a histidine kinase-like ATPbinding domain. The dimer axis runs perpendicular to the helices and

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Figure 6 Crystal structure of rat BCK. (A) Overall view of the rat BCK dimer, with the phospho-transfer reaction product ADP shown as a ball-and-stick model. Strands in the K domain are not labeled. (B) Same as (A), but rotated 90j around the horizontal line. (From Ref. 13.)

strands, which places the ATP-binding sites on opposite sides. The interface buries about 890 A˚2 of surface area per monomer and contains a number of hydrophobic and polar interactions, including the two ion pairs between Asp278 and His292 and between Arg286 and Asp365. BCK has a tendency to form tetramers in solution (26). A detailed analysis using analytical ultracentrifugation with sedimentation equilibrium showed that BCK exists in a dimer–tetramer equilibrium, with a dissociation constant Kd = 1.5 AM (data not shown). The BCK structures presented here do not shed light on the structural basis of tetramer formation; therefore, its physiological signiﬁcance remains unclear. B. The Presence of a Novel Structural K+ Ion in the Nucleotide-Binding Pocket Binding of adenosine nucleotides in BCK is uniquely mediated by both magnesium and potassium (Fig. 7). The magnesium ion is bound to the

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Figure 7 Nucleotide-binding pocket of rat BCK. Shown is superimposed 2Fo–Fc simulated annealing, with electron density maps (green, contoured at 1.5j) calculated for potassium (cyan) and magnesium (yellow) as well as bound ADP (A) and ATPgS (B). An anomalous diﬀerence Fourier electron density map (red, contoured at 5j) is shown for the ATPgS-bound BCK structure. The side chain of Phe303 has been removed for clarity. (From Ref. 13.) (See color insert.)

conserved N249 (N box) and the phosphate moieties of the nucleotide. The potassium ion is liganded by the g-phosphate of the nucleotide and carbonyl oxygens of Val298, Asp300, Phe303, and Gly337. No Electron density is observed in the potassium-binding site in apo-BCK, although high concentrations of KCl were present in the crystallization medium. The presence of potassium in the nucleotide-binding site explains the requirement of this metal ion for the kinase as well as ATPase activities of BCK (27). Rubidium, but not sodium, can substitute for potassium. Residues Val298 to Phe303 belong to a loop whose main-chain conformation is virtually identical to a potassium-binding loop in a helix–hairpin–helix motif in human DNA polymerase hV (28). Potassium stimulates the autophosphorylation activity of CheA (29), but CheA does not contain the potassium-binding motif present in BCK. On the other hand, the potassium-binding motif found in BCK is present in MutL. Interestingly, the sodium ion is a better ligand than potassium for nucleotide binding in MutL (W. Yang, personal communication). As described earlier, potassium-binding sites are also present in E1b, the primary substrate for BCK, although the motifs involved in binding potassium ions are diﬀerent between the two proteins.

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C. Nucleotide-Induced Domain Communication Nucleotide binding to the K domain of BCK induces structural changes that are transmitted via the B/K domain interface to the neighboring B domain (Fig. 8). In the apo-form, the region between Thr304 and Phe336 is disordered. Binding of ATPgS and potassium results in the ordering of residues Thr305–Ala306 and Gly335, which are adjacent to the potassium-binding site. This ordering leads to the formation of a quadruple aromatic stack comprising Tyr301, His302, Phe336, and Phe338. These residues alternate with potassium-binding residues, and their aromatic side chains are strategically placed in the interface between the B and K domains. The resultant aromatic stack contacts the adjacent B domain primarily through the Tyr301/ Asp161 and His302/Ala140 interactions, and causes a tilt in the top portion of the BH3/BH4 helix–hairpin–helix.

Figure 8 Superposition of the apo-structure of rat BCK (red) with its ADP-bound (blue) ATPgS-bound forms (green). Nucleotide (ADP or ATPgS) binding orders H302 and F336 (electron density not visible in the apo-form) in the B/K domain interface to form a quadruple aromatic stack between Y301, H302, F336, and F338. Structural diﬀerences in the BH3 and BH4 helices between the apo and the nucleotide-bound states are depicted. (From Ref. 13.) (See color insert.)

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The presence of the phospho-transfer reaction product ADP induces additional ordering of residues Pro332–Met333 in the vicinity of the nucleotide-binding site (Fig. 8). The carbonyl oxygen of His334 forms a hydrogen bond to the g-phosphate moiety, thereby eﬀectively trapping the ADP (Fig. 7). This trapping mechanism explains the characteristic product inhibition observed in mitochondral protein kinases (mPKs) (30) and renders ADP analogs attractive as potential inhibitors for these enzymes. Unlike typical protein kinases, BCK exhibits ATPase activity in the absence of its primary substrate, E1b. The product inhibition by ADP is likely to oﬀer a mechanism to prevent idle hydrolysis of ATP by BCK when its protein substrate E1b is absent. Reactivation of BCK requires the release of ADP, probably through speciﬁc removal of the loop region. These putative interactions could be provided by the LBD of E2b; lipoylated E2b has been shown to stimulate the kinase and ATPase activities of BCK (26,31). The formation and stabilization of the domain interface upon nucleotide binding could potentially provide an eﬀective mechanism for communication between the B and K domains. This structural coupling explains the eﬀects of certain site-speciﬁc mutations on enzyme activity. For example, replacing the central Tyr301 by alanine in the domain interface leads to a 95% reduction in the catalytic activity of BCK (26). A similar eﬀect was observed for the H121A mutation in Arabidopsis thaliana pyruvate dehydrogenase kinase (PDK) (32). This conserved histidine (His115 in human PDK1; His132 in rat BCK) is located 25 A˚ distant from the ATP binding site and is buried in the predominantly hydrophobic core of the B domain. Substitution by alanine creates a void that is expected to destabilize the core, thereby aﬀecting the structural integrity of this domain. Mutations that preserve the packing do not signiﬁcantly lower the activity, as observed for mutation H132N in BCK (data not shown). Thus, structural communication through the B/K domain interface may explain the long-range eﬀect of mutations in the core of the helical B domain on the nucleotide-binding pocket.

IV. SOLUTION STRUCTURE AND DYNAMICS OF THE LIPOYL-BEARING DOMAIN A. Function and Chain Fold of Human LBD The LBD of an a-ketoacid dehydrogenase complex mediates the transfer of the acyl moiety from the E1 to the E2 active site. After acyltransfer, the reduced lipoic acid moiety on LBD is reoxidized by E3, with NAD+ as the ultimate electron acceptor. Therefore, during the oxidative decarboxylation of a-ketoacids, LBD plays a central role in substrate channeling by visiting the three active sites in cognates E1b, E2b, and E3 (33). Moreover, in

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mammalian pyruvate dehydrogenase complex (PDC), the inner LBD mediates the feedback regulation of pyruvate dehydrogenase kinase activity by NAD+/NADH and CoA/acetyl-CoA. LBD plays this role by changing its oxidation, reduction, and acetylation state of the lipoyl group (34,35). Figure 9 shows the ribbon representation of the tertiary folding of the LBD of the human BCKD complex. Similar to other LBDs, the main feature of this structure is the presence of two h-sheets, which fold into the shape of a ﬂattened h-barrel. The order of h-strands in h-sheet S1 is h1-h8-h6-h3 and in h-sheet S2 h5-h4-h2-h7. The lipoylation-site residue Lys44 is located at the tip of a type I sharp turn of a h-hairpin formed by h4- and h5-strands on the S2 sheet. The N- and C-termini are close to each other in space and are located

Figure 9 Ribbon representation of the structure of human E2b LBD with lowest energy and no NOE violation. The structure consists of h-sheet S1 (h1, h3, h6, h8), and h-sheet S2 (h2, h4, h5, h7). The side-chain backbone of the lipoyl lysine residue, Lys-44, is shown. Each strand is numbered. L1 denotes the loop-connecting strands h1 and h2, and the N- and C-termini are labeled. (From Ref. 14.)

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on the far end of the S1 sheet. Another prominent feature of the structure is the presence of a V-shaped groove between the Lys44 h-hairpin and L1 loop connecting strands h1 and h2. The molecular packing in the middle of this groove is rather sparse, and a thin crevice f11 A˚ long is visible in this region in the space-ﬁlling model of LBD (structure not shown). There appears to be little contact between L1 and the Lys44 h-hairpin except for the charge interactions between the side chains of Lys44, Asp43, and Arg16. The interior core of the h-barrel is packed with hydrophobic residues, including Phe6, Val20, Trp23, Val31, Ile37, Ile49, Ile57, Leu60, Leu74, Ile77, and Leu82. B. Dynamics of Human LBD Reduced spectral density analysis of the relaxation data for human LBD discloses that most of the backbone amide groups of human LBD are rather rigid, with the exception of several mobile segments that are present in the solution structure. The dynamic behavior of the molecule can be visualized clearly from the sausage representations, which depict the sequence variation of J(H) (Fig. 10a) and J(O) (Fig. 10b). In both representations, the diameter of the sausage is related to the J-value of each residue and is adjusted to emphasize the diﬀerence in J-values between diﬀerent residues. For clarity, the diameters of the three C-terminal residues in Figure 10a were reduced.

Figure 10 Sausage representation of the sequence variation in the reduced spectral density functions J(522 MHZ) (a) and J(0) (b). The diameter of the sausage at a given location is related to the value of the reduced spectral density functions of the speciﬁc residue at that location. For clarity the diameters of the three carboxylterminal residues were purposely reduced, even though these residues are highly dynamic. (From Ref. 14.)

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Moreover, the radii of the sausage of internal residues whose amide protons cannot be observed, i.e., Glu13, Gly14, Arg16, and Asp43, were assigned the same values as the preceding residues. The distinct features of these structures are: (1) Residues in the lipoyl lysine-containing region comprising the L1 loop and the h-turn show picosecond-fast motion, as indicated by the large J(H) values; (2) slow millisecond-to-microsecond motion with large J(O) appears to be localized in and near the strands of S1 h-sheets, i.e., h1, h6, h7, and h8. Thus, the lipoyl Lys44 site of human LBD is the most ﬂexible region. In contrast, the S1 h-sheet, although rather rigid in the fast time scale, shows a slow conformational exchange motion. There is no indication of the presence of motion in the S2 h-sheet. The two strands forming the h-hairpin, i.e., h4 and h5, are also rather rigid. Perham and coworkers (36) have previously measured the 15N NMR relaxation parameters of LBD from E. coli PDC. Their results indicate that residues in and surrounding the lipoyl lysine h-turn are highly mobile and become less ﬂexible upon lipoylation of the lysine residue, which are in general in agreement with the present results. However, there are two major diﬀerences in the two LBDs. First, the L1 loop in the LBD of the human BCKD complex appears to be signiﬁcantly more mobile than the corresponding loop in E. coli LBD. Only residue Asp13 in the surface loop of E. coli LBD has NOE signiﬁcantly lower than residues in the bulk of the structure. Second, even though a slow conformational exchange motion can be clearly delineated in human LBD, no motion for h-sheet S1 was detected in E. coli LBD. The presence of high ﬂexibility in the lipoyl Lys44 site suggests that this lipoic acid–bearing site is not in a preformed conﬁguration. Binding of the ﬂexible lipoylation site region to its cognates E1b, E2b, E3 as well as BCK is likely to result in conformational rearrangements of interacting residues. This type of conformational adaptation presumably promotes optimal binding required for catalysis and regulation of the BCKD complex, similar to that described for E. coli thioesterase I (37) and dihydrofolate reductase (38). V. SUMMARY The past few years have witnessed the determination of structures for components and domain of the mammalian BCKD complex. The availability of this structural information constitutes an important impetus for understanding the structure–function relationship, protein–protein interactions, and the biochemical basis for disease states of the BCKD complex. Specifically, the human E1b structure shows a high degree of conservation at the cofactor-binding pocket in TDP-dependent enzymes. The structure further indicates that the prevalent Y393N-a MSUD mutation in the C-terminal tail, or the so-called ‘‘Mennonite region,’’ of the a-subunit disrupts a–hV and aV–h

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subunit interactions in E1b, resulting in the formation of inactive mutant heterodimers. Interestingly, novel potassium sites are present in both E1b and its regulatory enzyme BCK. Since the potassium ion concentration is high in mitochondria, it is plausible that both E1b and BCK utilize this monovalent metal ion for TDP and nucleotide binding as well as subunit assembly. The strict conservation in the nucleotide-binding K domain between BCK and GHL ATPases strongly suggests that the nucleotide-binding sites of proteins in this superfamily were evolved from a common ancestor. The nucleotideinduced formation of a hydrophobic stack at the interface between the B and K domains of BCK oﬀers a potential mechanism by which E2b stimulates the kinase activity. Finally, the solution structure for the LBD of the human BCKD complex depicts the signiﬁcant conservation of this domain among aketoacid dehydrogenase complexes. The high degree of ﬂexibility in the lipoyl-lysine region comprising the L1 loop and the Lys44 h-turn provides a structural basis for macromolecular interactions between the LBD domain and the E1b and BCK components of the mammalian BCKD complex.

ACKNOWLEDGMENTS This work was supported by grant DK-26758 from the National Institutes of Health and grant I-1286 from the Robert A. Welch Foundation. REFERENCES 1.

2.

3.

4.

5.

6.

FH Pettit, SJ Yeaman, LJ Reed. Puriﬁcation and characterization of branchedchain a-keto acid dehydrogenase complex of bovine kidney. Proc Natl Acad Sci USA 75:4881–4885, 1978. DT Chuang, VE Shih. Maple syrup urine disease (branched-chain ketoaciduria). The Metabolic and Molecular Basis of Inherited Disease. 8th ed. New York: McGraw-Hill, 2001, pp. 1971–2006 R Paxton, RA Harris. Isolation of rabbit liver branched chain a-ketoacid dehydrogenase and regulation by phosphorylation. J Biol Chem 257:14433– 144439, 1982. Z Damuni, ML Merryﬁeld, JS Humphreys, LJ Reed. Puriﬁcation and properties of branched-chain a-keto acid dehydrogenase phosphatase from bovine kidney. Proc Natl Acad Sci USA 81:4335–4338, 1984. RA Harris, JW Hawes, KM Popov, Y Zhao, Y Shimomura, J Sato, J Jaskiewicz, TD Hurley. Studies on the regulation of the mitochondrial a-ketoacid dehydrogenase complexes and their kinases. Adv Enzyme Regul 37:271–293, 1997. KG Cook, AP Bradford, SJ Yeaman, A Aitken, IM Fearnley, JE Walker. Regulation of bovine kidney branched-chain 2-oxoacid dehydrogenase complex by reversible phosphorylation. Eur J Biochem 145:587–591, 1984.

3D Structures of the Mammalian BCKD Complex 7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17. 18.

19.

20.

467

G Mattevi, E Obmolova, KH Schulze, AH Kalk, A Westphal, A de Kok, WG Hol. Atomic structure of the cubic core of the pyruvate dehydrogenase multienzyme complex. Science 255:1544–1550, 1992. A Ævarsson, K Seger, S Turley, JR Sokatch, WG Hol. Crystal structure of 2oxoisovalerate and dehydrogenase and the architecture of 2-oxo acid dehydrogenase multienzyme complexes. Nat Struct Biol 8:785–792, 1999. SS Mande, S Sarfaty, MD Allen, RN Perham, WG Hol. Protein–protein interactions in the pyruvate dehydrogenase multienzyme complex: dihydrolipoamide dehydrogenase complexed with the binding domain of dihydrolipoamide acetyltransferase. Structure 4:277–286, 1996. A Mattevi, G Obmolova, JR Sokatch, C Betzel, WG Hol. The reﬁned crystal structure of Pseudomonas putida lipoamide dehydrogenase complexed with NAD+ at 2.45-A˚ resolution. Proteins 4:336–351, 1992. JE Knapp, D Carroll, JE Lawson, SR Ernst, LJ Reed, ML Hackert. Expression, puriﬁcation, and structural analysis of the trimeric form of the catalytic domain of the Escherichia coli dihydrolipoamide succinyltransferase. Protein Sci 9:37– 48, 2000. A Ævarsson, JL Chuang, RM Wynn, S Turley, DT Chuang, WG Hol. Crystal structure of human branched-chain a-ketoacid dehydrogenase and the molecular basis of multienzyme complex deﬁciency in maple syrup urine disease. Structure 8:277–291, 2000. M Machius, JL Chuang, RM Wynn, DR Tomchick, DT Chuang. Structure of rat BCKD kinase: nucleotide-induced domain communication in a mitochondrial protein kinase. Proc Natl Acad Sci USA 98:11218–11223, 2001. CF Chang, HT Chou, JL Chuang, DT Chuang, TH Huang. Solution structure and dynamics of the lipoic acid–bearing domain of human mitochondrial branched-chain a-keto acid dehydrogenase complex. J Biol Chem 277:15865– 15873, 2002. RM Wynn, JR Davie, JL Chuang, CD Cote, DT Chuang. Impaired assembly of E1 decarboxylase of the branched-chain a-ketoacid dehydrogenase complex in type IA maple syrup urine disease. J Biol Chem 273:13110–13118, 1998. Y Lindqvist, G Schneider, U Ermler, M Sundstrom. Three-dimensional structure of transketolase, a thiamine diphosphate–dependent enzyme, at 2.5-A˚ resolution. EMBO J 11:2373–2379, 1992. 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. 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. 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. E Chabrie`re, MH Charon, A Volbeda, L Pieulle, EC Hatchikian, C Fontecilla-

468

21. 22.

23.

24. 25. 26.

27.

28.

29. 30.

31.

32.

33.

34.

Chuang et al. Camps. Crystal structures of the key anaerobic enzyme pyruvate: ferredoxin oxidoreductase, free and in complex with pyruvate. Nat Struct Biol 6:182–190, 1999. CF Hawkins, A Borges, RN Perham. A common structural motif in thiamine pyrophosphate–binding enzymes. FEBS Lett 255:77–82, 1989. CR Fisher, CW Fisher, DT Chuang, RP Cox. Occurrence of a Tyr393!Asn (Y393N) mutation in the E1a gene of the branched-chain a-keto acid dehydrogenase complex in maple syrup urine disease patients from a Mennonite population. Am J Hum Genet 49:429–434, 1991. I Matsuda, Y Nobukuni, H Mitsubuchi, Y Indo, F Endo, J Asaka, H Harad. A T!A substitution in the E1a subunit gene of the branched-chain a-ketoacid dehydrogenase complex in two cell lines derived from Mennonite maple syrup urine disease patients. Biochem Biophys Res Commun 172:451–464, 1990. L Marshall, A DiGeorge. Maple syrup urine disease in the old-order Mennonites. Am J Hum Genet 33:139, 1981. DT Chuang. Maple syrup urine disease: it has come a long way. J Pediatr 132:S17–S23, 1998. RM Wynn, JL Chuang, CD Cote, DT Chuang. Tetrameric assembly and conservation in the ATP-binding domain of rat branched-chain a-ketoacid dehydrogenase kinase. J Biol Chem 275:30512–30519, 2000. Y Shimomura, MJ Kuntz, M Suzuki, T Ozawa, RA Harris. Monovalent cations and inorganic phosphate alter branched-chain a-ketoacid dehydrogenase kinase activity and inhibitor sensitiviy. Arch Biochem Biophys 266:210–218, 1988. H Pelletier, MR Sawaya. Characterization of the metal ion–binding helix– hairpin-helix motifs in human DNA polymerase h by X-ray structural analysis. Biochemistry 35:12778–12787, 1996. JF Hess, K Oosawa, N Kaplan, MI Simon. Phosphorylation of three proteins in the signaling pathway of bacterial chemotaxis. Cell 53:79–87, 1988. TE Roche, LJ Reed. Monovalent cation requirement for ADP inhibition of pyruvate dehydrogenase kinase. Biochem Biophys Res Commun 59:1341–1348, 1974. JR Davie, RM Wynn, M Meng, YS Huang, G Aalund, DT Chuang, KS Lau. Expression and characterization of branched-chain a–ketoacid dehydrogenase kinase from the rat. Is it a histidine-protein kinase? J Biol Chem 270:19861– 19867, 1995. JJ Thelen, JA Miernyk, DD Randall. Pyruvate dehydrogenase kinase from Arabidopsis thaliana: a protein histidine kinase that phosphorylates serine residues. Biochem J 349:195–201, 2000. RN Perham. Swing arms and swinging domains in multifunctional enzymes: Catalytic machines for multistep reactions. Annu Rev Biochem 69:961–1004, 2000. GA Radke, K Ono, S Ravindran, TE Roche. Critical role of a lipoyl cofactor of the dihydrolipoyl acetyltransferase in the binding and enhanced function of the pyruvate dehydrogenase kinase. Biochem Biophys Res Commun 1993:982–991, 190.

3D Structures of the Mammalian BCKD Complex

469

35. JC Baker, X Yan, T Peng, S Kasten, TE Roche. Marked diﬀerences between two isoforms of human pyruvate dehydorgenase kinase. J Biol Chem 275:15773– 15781, 2000. 36. 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. 37. YT Yuang, YC Liaw, VY Gorbatyuk, TH Huang. Backbone dynamics of Escherichia coli thioesterase/protease I: evidence of a ﬂexible active-site environment for a serine protease. J Mol Biol 307:1075–1090, 2001. 38. GP Miller, SJ Benkovic. Stretching exercises—ﬂexibility in dihydrofolate reductase catalysis. Chem Biol 5:R105–R113, 1998.

26 Variability of Human Pyruvate Dehydrogenase Complex Deficiency Douglas S. Kerr and Christine L. Schmotzer Case Western Reserve University School of Medicine, Cleveland, Ohio, U.S.A.

I. INTRODUCTION Defects of the human pyruvate dehydrogenase complex (PDC) may cause potentially severe neurological disease or death, but they are present with great variability. In recent years, much has been learned about the molecular basis of these disorders, which predominantly aﬀect the E1a subunit of pyruvate dehydrogenase (1,2). However, factors that account for the great variability observed in aﬀected individuals have been only partially explained. A better understanding of these variables may provide future guidance to ﬁnding more eﬀective therapy, which at present is of limited beneﬁt. Pyruvate dehydrogenase complex deﬁciency is one of a number of disorders of mitochondrial function resulting in impaired production of ATP, accumulation of lactic acid, and typically severe systemic dysfunction. Amongst these disorders, it is estimated that PDC deﬁciency accounts for approximately 10%, which makes it one of the more commonly diagnosed causes of defective mitochondrial energy production. These disorders diﬀer from one another due to the tissue speciﬁcity of metabolic fuel requirements. Pyruvate dehydrogenase complex deﬁciency primarily aﬀects the nervous 471

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system and has little pathological eﬀect on the heart, skeletal muscle, or other organ function. This is consistent with the dependence of the central nervous system on glucose, even in the fasting state. By contrast, defects of fatty acid oxidation have major eﬀects on the heart, skeletal muscle, and frequently liver, which are tissues that oxidize fatty acids directly in both the fed and fasting states. Although disorders of fatty acid oxidation may have severe consequences for brain function, these consequences are typically mediated through hypoglycemia resulting from inability to derive suﬃcient energy by fatty acid oxidation while fasting. Defects of the electron transport chain aﬀect both pyruvate oxidation and fatty acid oxidation and typically have systemic consequences aﬀecting the central nervous system, heart, skeletal muscle, and in many cases other organs and tissues. II. CLINICAL, METABOLIC, AND PATHOLOGICAL CONSEQUENCES PDC deﬁciency is typically associated with the accumulation of lactate, pyruvate, and alanine in body ﬂuids (2). The amounts of these increases may be quite variable, from barely detectable to severe metabolic acidosis. However, in almost all cases the ratio of blood lactate to pyruvate (reﬂecting the intracellular ratio of NADH to NAD) is within the normal range of 10 to 20. This is in distinction to defects of electron transport chain, where the ratio of lactate to pyruvate is predictably high. Unfortunately, accurate measurements of blood pyruvate are not readily available in many clinical laboratories. In some cases, elevated lactate and pyruvate is limited to spinal ﬂuid or to the brain, where lactate is potentially detectable by magnetic resonance spectroscopy. Lactate and pyruvate concentrations are typically increased following a carbohydrate meal. This is in contrast to defects of gluconeogenesis, including pyruvate carboxylase deﬁciency, in which lactate increases during fasting, frequently associated with hypoglycemia. In contrast to defects of fatty acid oxidation of the electron transport chain, PDC deﬁciency is not typically associated with the accumulation of intermediates of the Krebs cycle or incomplete fatty acid oxidation. The clinical presentation of PDC deﬁciency is not distinct from many other disorders of impaired mental development or mitochondrial dysfunction. The frequency and severity of these manifestations is remarkably variable, so a common phenotype does not diagnostically characterize the majority of individuals with this disorder. In a preliminary review of medical records of approximately 130 cases of conﬁrmed PDC deﬁciency evaluated in our laboratory, we found the most common clinical problem to be developmental delay (>75%), frequently associated with hypotonia (57%) (Table1).

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Table 1 Frequency of Reported Features of PDC Deﬁciency Feature

% Aﬀected (n = 131)

Developmental delay Hypotonia Seizure disorder Apnea, hypoventilation Ataxia Microcephaly Dysmorphic features Poor growth Peripheral neuropathy CNS degenerationa Congenital brain malformation

>75 57 38 36 18 15 13 11 3 <10 <5

a

Including Leigh syndrome. Source: DS Kerr and C Schmotzer, unpublished data.

Seizures are also relatively common (38%). Less common but more distinctive clinical features include ataxia (18%), peripheral neuropathy, apnea and central respiratory failure, dysmorphic features, congenital anomalies of brain development (especially absence of the corpus callosum), and Leigh syndrome (degeneration of the basal ganglia and brain stem). Neuropathological ﬁndings in cases that have died and had postmortem examination or that have had accurate brain imaging include microcephaly (indicating impaired brain growth), absence or hypoplasia of the corpus callosum (the most common congenital anomaly), and degeneration of the basal ganglia and brainstem. There have been surprisingly few other systemic pathological ﬁndings. A more thorough review of these cases is in progress. Diagnosis of PDC deﬁciency typically follows clinical suspicion and detection of lactic acidemia. Assays most often are done in cultured skin ﬁbroblasts based on thiamine pyrophosphate–dependent release of 14CO2 derived from 1-14C-pyruvate (3). This is usually done after activation of PDC by preincubation with dichloroacetate (intact cells) or pyruvate dehydrogenase phosphatase (frozen tissues). We have found that assay of PDC in blood lymphocytes is a useful adjunct for detection of PDC deﬁciency, since blood can be more readily obtained and several weeks are not required for ﬁbroblast culture (4). Furthermore, ﬁnding normal PDC activity in cells or tissues does not exclude the diagnosis, since PDC deﬁciency may be variably expressed in cells and tissues, not only in aﬀected females but also in aﬀected males, for

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unexplained reasons (4). Therefore, reliable diagnosis or diagnostic conﬁrmation typically requires assays in two or more specimen types. If low activity is found, follow-up assays are typically performed of the activity E1, E2, and E3 catalytic subcomponents and/or immunoassays of these components. The latter may be especially useful in the detection of heterozygous females with E1 subunit deﬁciency resulting in somatic cell mosaicism. III. GENETIC BASIS OF PYRUVATE DEHYDROGENASE DEFICIENCY The vast majority (greater than 90%) of proven mutations causing decreased activity of PDC have been found within the coding region gene for the E1a subunit. This gene, located on chromosome X (Xp22.3), includes 11 exons and encodes a 41-kDa protein that forms a heteroduplex with the E1h subunit (5). A recent multicenter review reported 76 diﬀerent mutations occurring in 130 subjects, including 37 missense/nonsense point mutations and 39 insertions and deletions; the latter usually result in the synthesis of an incomplete, unstable peptide (6). No ‘‘null’’ large deletions have been described in males, but some exon-skipping mutations have been described in females. Because of random patterns of inactivation of the two alleles on chromosome X in females, a range of expression of E1a is expected, accounting for the ability of females to survive with more severe mutations (7). This may also contribute to underestimation of the frequency of E1a deﬁciency in heterozygous females. The prevalence of E1a deﬁciency among siblings of identiﬁed aﬀected subjects is less than 10%, implying a relatively high rate of new mutations and/or embryonic and fetal loss postfertilization (8). A second, processed (intronless) gene for the E1a subunit is located on chromosome 4 and is uniquely expressed in developing spermatozoa (9). Since carbohydrate oxidation is essential to ATP production in the sperm, expression of this alternative gene accounts for the viability of mature sperm despite the absence or normal inactivation of chromosome X. An analogous situation exists for phosphoglycerokinase. The mechanism of regulation of expression of this second gene has been investigated in the mouse and shown to depend on demethylation of critical regions of the promoter as well as on testisspeciﬁc transcription factors (10). A truncated, demethylated construct of this promoter can be expressed in somatic cells. To date, no mutations of the E1h subunit have been reported in human PDC deﬁciency. Mutations of E1a resulting in instability and loss of immunoreactive protein are commonly associated with concomitant loss of both E1a and h immunoreactivity, presumably reﬂecting the critical role of binding between the a and h subunits for their mutual stability. Mutations have been

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described in the less common defects aﬀecting E2, E3 binding protein and E3 (11). E3 deﬁciency results in multiple a-ketoacid dehydrogenase deﬁciencies (pyruvate, a-ketoglutarate, and branch-chain a-ketoacid dehydrogenase complexes). Defects of pyruvate dehydrogenase phosphatase result in failure to activate the complex. IV. VARIABILITY OF OUTCOME OF PDC DEFICIENCY To what extent can the variability of clinical manifestations be attributed to known molecular mechanisms of PDC deﬁciency? Some of these factors are indeed predictable. Most notable is the variable pattern of inactivation of the two alleles in heterozygous females with E1a deﬁciency (7). Furthermore, there is a large heterogeneity of E1a mutations, and each of these could be expected to have a somewhat diﬀerent consequence for PDC activity. At least one of the described E1a exon-skipping mutations has been shown to be result in only partial exon skipping, accounting for its occurrence in surviving males (12). However, these considerations are not suﬃcient to explain all clinical observations. Males with the most common E1a point mutation (R234G) have had quite diﬀerent outcomes in terms of in vitro activity, survival, and mental development (Table 2). This mutation was initially discovered in male brothers who had normal activity of PDC in cultured skin ﬁbroblasts but deﬁciency in lymphocytes and other tissues (13). However, some males with the same mutation have lower activity in the ﬁbroblasts (Table 2). It should be noted that ﬁbroblast PDC activity does not correlate well with long-term survival. Some of the oldest, highestfunctioning males with E1a deﬁciency have barely detectable activity in their skin ﬁbroblasts (3). Presumably, other environmental factors may play a role, including nutritional factors (see later). But to date, correlation with these variables has been quite limited. We speculated that, conceivably, expression of the E1a gene on chromosome 4 (Pdha2) in somatic cells might contribute to some of this unaccounted variation. To investigate this possibility, we assayed expression of the Pdha2 gene in several skin ﬁbroblast lines from males with the R234G mutation that showed a range of PDC activity. We found no detectable message from the Pdha2 gene and no diﬀerence in the level of mRNA from the Pdha1 (chromosome X) gene, estimated by an RNAse protection assay (14). This hypothetical possibility cannot be excluded in other situations. Since the observed diﬀerences of activity of PDC associated with this mutation correlate with levels of immunoreactivity of E1a and h, it is assumed that whatever variables may contribute to this diﬀerence aﬀect the stability of a tenuous association of the two subunits.

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Table 2 Clinical and Biochemical Variability of PDC-Deﬁcient Cases Due to the E1a R234G Mutation Sex

Age (yr)b

Aliveb

Family relation

PDC activity % (sample)c

Mental development

1 2 3

M M F

3 9 8

No No Yes

Brother Brother Sister

77.(f ), 4 (l) 168.(f ), 7 (l) 53.(f ), 110 (l)

Delayed Delayed Mild delay

4 5 6 7 8 9 10

F M F M F M M

Adult 9 Adult 8 Adult 7 10

Yes Yes Yes Yes Yes Yes Yes

Mother Son Mother Son Mother

112.(f ), 35 (l) 39.(f ) 40.(f ) 48.(f ) 40.(f ) 16.(f ) 40.(f )

Normal Delayed Delayed Delayed Delayed Delayed Normal

11

M

12

M

12

No

13

M

16

No

14

M

4

Yes

15

M

4

No

16

M

6

17 18

F M

19

M

Casea

50.

Delayed

Brother

36.(f )

Delayed

Brother

32.(f )

Normal

31.(f )

Delayed

Brother

40.(l)

Delayed

No

Brother

10.(m)

Delayed

Adult 2

Yes No

Mother

77.(l) 46.(f ), 17 (l)

Normal Delayed

14

Yes

19.(m)

Delayed

Other features Hypotonia Hypotonia Hypotonia, ataxia Ataxia Ataxia Ataxia Ataxia Ataxia, neuropathy Leigh syndrome Hypotonia, ataxia Hypotonia, ataxia Leigh syndrome Thiamin responsive Leigh syndrome Hypotonia, Leigh syndrome Hypotonia, Leigh syndrome

a

Cases are grouped as families. Status at time of publication or last update. c Sample type: f = ﬁbroblasts, l = lymphocytes/blasts, m = muscle. b

V. TREATMENT Unfortunately, no general, very eﬀective treatment has been described for PDC deﬁciency. Several strategies have been employed, with variable success. These include the use of a ketogenic diet, supplementation with thiamine, and administration of dichloroacetate (to inhibit PDH kinase). For the future, gene therapy is possible. Ketogenic diet therapy is based on a sound rationale,

Ref. 4., 13 4., 13 6. 4., 13 33. 34. 34. 34. 34. 6. 35. 16. 16. 36. 18. 18. 18. 37.

37.

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477

providing a source of energy and acetyl-CoA from fatty acid oxidation while minimizing production of pyruvate from glycolysis and stimulating ketosis, providing an alternate to glucose for brain utilizaton. Initial reports in single cases treated with ketogenic diets seemed promising (15). We reviewed the outcome of seven males with known mutations of E1a, three with a R349H mutation and four with the more common R234G mutation (two pairs of brothers) (16). The conclusion was that the more severely carbohydrate was restricted in the diet (down to less than 7% of total energy) and the earlier such a diet was implemented, the longer the aﬀected children survived and the greater their mental development. However, all of these children died at a relatively young age, so although ketogenic diet therapy may be beneﬁcial, it clearly is not a suﬃcient solution. It also is very diﬃcult to sustain ketosis over a long period of time, but some families have been remarkably successful in doing so, with apparently good results. Thiamine in variable amounts well above the recommended dietary allowance has commonly been given to individuals with PDC deﬁciency. Over 10 cases have been reported in which it was claimed that administration of thiamine improved the clinical outcome and/or reduced blood lactate (selected cases are summarized in Table 3). Some of these cases have been treated with gram quantities of thiamine (over a thousand times above the RDA). There has been no controlled trial to evaluate the eﬃcacy of thiamine therapy for PDC deﬁciency. Cases with certain E1a mutations may be particularly susceptible to this reported beneﬁcial eﬀect (17–19). In vitro studies with cultured lymphoblasts showed substantial diﬀerences in the concentration dependence of thiamine pyrophosphate required for PDC activity, suggesting a diﬀerence in TPP binding to pyruvate dehydrogenase. There is a discrepancy between these observations and other observations of the kinetics of PDC dependence on TPP. The concentration of TPP required for half-maximum activity reported in normal lymphoblast homogenates (apparent ‘‘Ka’’) is much lower (<109 M) than the Ka reported for puriﬁed recombinant human PDC (7 108 M) or puriﬁed pig heart PDC (2 107 M ), and the reported apparent ‘‘Ka’’ for lymphoblasts with an E1a H44R point mutation (approx. 5 105 M) is much higher than found with this same recombinant mutant PDC (2 x 107 M ) (20,21). Recombinant PDC with the more common R234G mutation, which has also been reported to be thiamine responsive, shows a sevenfold increase in the Ka for TPP (22). However, the concentration of TPP is increased less than twofold in liver mitochondria from rats with a 10-fold higher than normal intake of thiamine (23). TPP is involved in the binding of E1a and h, and addition of TPP protects recombinant normal E1 from heat inactivation. But the concentration dependence of this protection is not diﬀerent in the H44R mutation (20). Therefore, it is still not clear by exactly

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Table 3 Thiamine-Responsive Cases of PDC Deﬁciency PDC activity % (sample)b

Case

Sex

Age (yr)a

1

M

6

12 (f )

2

M

3

44 (f )

3

M

8

36 (f )

4

M

8

5

M

1

17 (f )

6

M

9

25 (f )

7

M

4

28 (m)

8

M

6

43 (l)

a b

8 (m)

Clinical features Optic atrophy, ataxia Hypotonia, ataxia, delayed Hypotonia, ataxia Neuropathy Hypotonia, poor growth Exercise intolerance Hypotonia, myoclonus Delayed, Leigh syndrome, died

Dose of thiamine (mg/d) 600

Reported response

PDC defect

Ref.

Improved

Unknown

38

1800

Much improved

Unknown

39

600

Much improved Much improved Improved

E1a N297ins

40

E1

41

E1a H15R

17

Improved

E1a V360del

42

Much improved Improved

Unknown

43

E1aR234G

18

500 600 1000 900 500

Status at time of publication. Sample type: f = skin ﬁbroblasts, m = muscle biopsy, lymphoblasts (assayed with low TPP).

what mechanism clinical responsiveness to thiamine correlates with these in vitro observations. There have been a number of anecdotal reports that administration of dichloroacetate has been beneﬁcial to children with PDC deﬁciency, but these were short-term studies and a favorable outcome was largely based on lowering of blood lactate (24). Recently, a controlled clinical trial of the potential beneﬁts of DCA treatment for children with congenital lactic acidosis has been conducted at the University of Florida. Initial analysis of these results has indicated no diﬀerence in the outcome of the children with PDC deﬁciency while they were receiving or not receiving DCA (25). Although it initially was not clear exactly how inhibition of pyruvate dehydrogenase kinase by DCA could increase activation of a genetically altered enzyme, in vitro studies have shown that DCA may stabilize the E1a component in certain mutations (26). Long-term hopes for eﬀective gene therapy are based on very preliminary experiments in vitro and development of promising animal models. Pyruvate dehydrogenase activity can be increased by transfecting E1-deﬁcient

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human ﬁbroblasts with an adeno-associated virus vector construct (27) and also by transfected cDNA (28–30). Further studies are necessary to determine conditions for stable transfection and for expression in the central nervous system. Mouse models have now been developed for both E1a deﬁciency as well as E3 deﬁciency (31,32). Absence of E1a or E3 results in interruption of fetal development. Hypomorphic variations of these models should provide useful opportunities for further investigation of gene therapy or combinations of gene therapy with dietary treatment and possible supplementation with thiamine or DCA.

VI. CONCLUSIONS Analysis of PDC deﬁciency in both humans and experimental animals has shown that some residual activity of the enzyme is required for intrauterine development and extrauterine survival. Exactly how much is required is uncertain, but it is probable that small increments in activity could have signiﬁcant clinical beneﬁts. Variations of activity and clinical outcomes are to a large extent explained by the large variety of types of mutations and by the X-linked location of the gene for E1a causing variable expression in aﬀected females. However, there is unexplained signiﬁcant variation in aﬀected males with regard to both enzyme activity in vitro as well as clinical outcome. Other genetic, nutritional, and environmental factors may contribute to such variation. Identiﬁcation of those variables could provide clues to better therapeutic strategies. None of the current therapies are extremely eﬀective, although there is a rationale for expecting that diﬀerent mutations would respond more or less to speciﬁc interventions, such as administration of thiamine or dichloroacetate. Combinations of these with ketogenic diet treatment may be most eﬀective for now. Applications of gene therapy, especially in mouse models, hold promise for the future intervention, provided that stable expression can be achieved within the central nervous system. Eﬀective application of potential new therapies to human disease requires well-controlled clinical trials, which are particularly expensive and diﬃcult to organize for rare diseases.

ABBREVIATIONS PDC E1a, E1h E2 E3 TPP

pyruvate dehydrogenase complex alpha and beta subunits of the pyruvate dehydrogenase component dihydrolipoamide transacetylase dihydrolipoamide dehydrogenase thiamine pyrophosphate

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REFERENCES 1.

2.

3. 4.

5.

6.

7. 8.

9.

10.

11.

12.

13.

14.

BH Robinson. Lactic acidemia (Disorders of pyruvate carboxylase, pyruvate dehydrogenase). In: CR Scriver, AL Beaudet, WS Sly, eds. Metabolic and Molecular Basis of Inherited Disease. 7th ed. New York: McGraw-Hill, 1995, pp 1479–1499. DS Kerr, ID Wexler, AB Zinn. The pyruvate dehydrogenase complex and tricarboxylic acid cycle. In: J Fernandes, J-M Saudubray, eds. Inherited Metabolic Diseases. Diagnosis and Treatment. 3rd ed. Berlin: Springer-Verlag, 2000, pp. 127–138. KFR Sheu, CWC Hu, MF Utter. Pyruvate dehydrogenase complex activity in normal and deﬁcient ﬁbroblasts. J Clin Invest 67:1463–1471, 1981. DS Kerr, SA Berry, MM Lusk, L Ho, MS Patel. A deﬁciency of both subunits of pyruvate dehydrogenase which is not expressed in ﬁbroblasts. Pediatr Res 24: 95–100, 1988. RM Brown, HHM Dahl, GK Brown. X-chromosome localization of the functional gene for the E1a subunit of the human pyruvate dehydrogenase complex. Genomics 4:174–181, 1989. W Lissens, L De Meirleir, S Seneca, I Liebaers, GK Brown, RM Brown, M Ito, E Naito, Y Kuroda, DS Kerr, ID Wexler, MS Patel, BH Robinson, A Seyda. Mutations in the X-linked pyruvate dehydrogenase (E1) alpha subunit gene (PDHA1) in patients with a pyruvate dehydrogenase complex deﬁciency. Hum Mutat 15:209–219, 2000. HHM Dahl. Pyruvate dehydrogenase E1a deﬁciency: Males and females diﬀer yet again. Am J Hum Genet 56:553–557, 1995. DS Kerr, MM Lusk. Infrequent expression of heterozygosity or deﬁciency of pyruvate dehydrogenase (E1) among parents and sibs of aﬀected patients. Pediatr Res 31:133A, 1992. HHM Dahl, RM Brown, WM Hutchison, C Maragos, GK Brown. A testisspeciﬁc form of the human pyruvate dehydrogenase E1 alpha subunit is coded for by an intronless gene on chromosome 4. Genomics 8:225–232, 1990. RC Iannello, J Young, S Sumarsono, MJ Tymms, HH Dahl, J Gould, M Hedger, I Kola. Regulation of Pdha-2 expression is mediated by proximal promoter sequences and CpG methylation. Mol Cell Biol 17:612–619, 1997. DS Kerr, ID Wexler, A Tripatara, MS Patel. Defects of the human pyruvate dehydrogenase complex. In: MS Patel, T Roche, eds. Alpha Keto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996, pp 249–270. AK Cardozo, L De Meirleir, I Liebaers, W Lissens. Analysis of exonic mutations leading to exon skipping in patients with pyruvate dehydrogenase E1 alpha deﬁciency. Pediatr Res 48:748–753, 2000. ID Wexler, SG Hemalatha, TC Liu, SA Berry, DS Kerr, MS Patel. A mutation in the E1a subunit of pyruvate dehydrogenase associated with variable expression of pyruvate dehydrogenase complex deﬁciency. Pediatr Res 32:169–174, 1992. DS Kerr, Rice E, Korytko A, Chen Y, Y Du, M Lusk, MS Patel, ID Wexler. Expression of the testis-speciﬁc pyruvate dehydrogenase E1a subunit in normal

Variability of Human PDC Deficiency

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

481

and E1a deﬁcient human cells and tissues. Annual Meeting of the Society for Inherited Metabolic Disorders, 1998. RE Falk, SD Cederbaum, JP Blass, GE Gibson, RA Pieter Kark, RE Carrel. Ketogenic diet in the management of pyruvate dehydrogenase deﬁciency. Pediatrics 58:713–721, 1976. ID Wexler, SG Hemalatha, J McConnell, NR Buist, HH Dahl, SA Berry, SD Cederbaum, MS Patel, DS Kerr. Outcome of pyruvate dehydrogenase deﬁciency treated with ketogenic diets. Studies in patients with identical mutations. Neurology 49:1655–1661, 1997. E Naito, M Ito, E Takeda, I Yokota, S Yoshijima, Y Kuroda. Molecular analysis of abnormal pyruvate dehydrogenase in a patient with thiamine-responsive congenital lactic acidemia. Pediatr Res 36:340–346, 1994. E Naito, M Ito, I Yokota, T Saijo, J Matsuda, H Osaka, S Kimura, Y Kuroda. Biochemical and molecular analysis of an X-linked case of Leigh syndrome associated with thiamin-responsive pyruvate dehydrogenase deﬁciency. J Inherit Metab Dis 20:539–548, 1997. E Naito, M Ito, I Yokota, T Saijo, Y Ogawa, Y Kuroda. Diagnosis and molecular analysis of three male patients with thiamine-responsive pyruvate dehydrogenase complex deﬁciency. J Neurol Sci 201:33, 2002. SJ Jacobia, LG Korotchkina, MS Patel. Characterization of a missense mutation at histidine-44 in a pyruvate dehydrogenase–deﬁcient patient. Biochim Biophys Acta 1586:32–42, 2002. S Strumilo, J Czemiecki, P Dobrzyn. Regulatory eﬀect of thiamin pyrophosphate on pig heart pyruvate dehydrogenase complex. Biochem Biophys Res Commun 256:341–345, 1999. SJ Jacobia, LG Korotchkina, MS Patel. Diﬀerential eﬀects of two mutations at arginine-234 in the alpha subunit of human pyruvate dehydrogenase. Arch Biochem Biophys 395:121–128, 2001. PV Blair, R Kobayashi, HM Edwards, NF Shay, DH Baker, RA Harris. Dietary thiamin level inﬂuences levels of its diphosphate form and thiamin-dependent enzymic activities of rat liver. J Nutr 129:641–648, 1999. PW Stacpoole, CL Barnes, MD Hurbanis, SL Cannon, DS Kerr. Treatment of congenital lactic acidosis with dichloroacetate: a review. Arch Pediatr Adolesc Med 77:535–541, 1997. PW Stacpoole. Dichloroacetate treatment of congenital lactic acidosis: preliminary outcome results of the DCA/CLA trial. Annual Meeting of the Society for Inherited Metabolic Disorders, p. 11, 2002. KJ Morten, P Beattie, GK Brown, PM Matthews. Dichloroacetate stabilizes the mutant E1-alpha subunit in pyruvate dehydrogenase deﬁciency. Neurology 53:612–616, 1999. R Owen, RJ Mandel, CV Ammini, TJ Conlon, DS Kerr, PW Stacpoole, TR Flotte. Gene therapy for pyruvate dehydrogenase E1-alpha deﬁciency using recombinant adeno-associated virus 2 (rAAV2) vectors. Mol Ther 6:394–399, 2002.

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28. T Saijo, E Naito, M Ito, I Yokota, J Matsuda, Y Kuroda. Stable restoration of pyruvate dehydrogenase complex in E1-defective human lymphoblastoid cells: evidence that three C-terminal amino acids of E1-alpha are essential for the structural integrity of heterotetrameric E1. Biochem Biophys Res Commun 228:446–451, 1996. 29. K Chun, BH Robinson. Expression of normal and mutant pyruvate dehydrogenase complex E1-alpha cDNAs in cultured human lymphoblasts. Arch Biochem Biophys 349:246–250, 1998. 30. A Seyda, BH Robinson. Functional expression of four PDH-E(1)alpha recombinant histidine mutants in a human ﬁbroblast cell line with zero endogenous PDH complex activity. Biochem Biophys Res Commun 270:1068– 1073, 2000. 31. MT Johnson, HS Yang, T Magnuson, MS Patel. Targeted disruption of the murine dihydrolipoamide dehydrogenase gene (Dld) results in perigastrulation lethality. Proc Natl Acad Sci USA 94:14512–14517, 1997. 32. MT Johnson, S Mahmood, SL Hyatt, HS Yang, PD Soloway, RW Hanson, MS Patel. Inactivation of the murine pyruvate dehydrogenase (Pdha 1) gene and its eﬀect on early embryonic development. Mol Genet Metab 74:293–302, 2001. 33. K Chun, N MacKay, R Petrova-Benedict, BH Robinson. Mutations in the Xlinked E1a subunit of pyruvate dehydrogenase leading to deﬁciency of the pyruvate dehydrogenase complex. Hum Mol Genet 2:449–454, 1993. 34. K Chun, N MacKay, R Petrova-Benedict, A Federico, A Fois, DEC Cole, E Robertson, BH Robinson. Mutations in the X-linked E1a subunit of pyruvate dehydrogenase: exon skipping, insertion of duplicate sequence, and missense mutations leading to the deﬁciency of the pyruvate dehydrogenase complex. Am J Hum Genet 56:558–569, 1995. 35. P Briones, MJ Lopez, L De Meirleir, A Ribes, M Rodes, C Martinez-Costa, M Peris, W Lissens. Leigh syndrome due to pyruvate dehydrogenase E1-alpha deﬁciency (point mutation R263G) in a Spanish boy. J Inherit Metab Dis 19: 795–796, 1996. 36. W Lissens, L De Meirleir, S Seneca, C Benelli, C Marsac, BT Poll-The, P Briones, W Ruitenbeek, O van Diggelen, D Chaigne, V Ramaekers, I Liebaers. Mutation analysis of the pyruvate dehydrogenase E1-alpha gene in eight patients with a pyruvate dehydrogenase complex deﬁciency. Hum Mutat 7:46–51, 1996. 37. C Marsac, C Benelli, I Desguerre, M Diry, F Fouque, L De Meirleir, G Ponsot, S Seneca, F Poggi, JM Saudubray, MT Zabot, D Fontan, W Lissens. Biochemical and genetic studies of four patients with pyruvate dehydrogenase E1alpha deﬁciency. Hum Genet 99:785–792, 1997. 38. JP Blass, D Lonsdale, BW Uhlendorf, E Ham. Intermittent ataxia with pyruvate decarboxylase activity. Lancet 1:1302, 1971. 39. H Wick, K Schweizer, R Baumgartner. Thiamine dependency in a patient with congenital lactic acidaemia due to pyruvate dehydrogenase deﬁciency. Agents Actions 7:405–410, 1977. 40. T Fujii, MBG Alvarez, KFR Sheu, PJ Kranzeble, DC DeVivo. Pyruvate

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dehydrogenase deﬁciency: the relation of the E1-alpha mutation to the E1-beta subunit deﬁciency. Pediat Neurol 14:328–334, 1996. 41. G Bonne, C Benelli, L De Meirleir, W Lissens, M Chaussain, M Diry, JP Clot, G Ponsot, V Geoﬀroy, JP Leroux, et al. E1 pyruvate dehydrogenase deﬁciency in a child with motor neuropathy. Pediatr Res 33:284–288, 1993. 42. K Narisawa, H Endo, S Miyabayashi, K Tada. Thiamine responsive pyruvate dehydrogenase deﬁciency. J Nutr Sci Vitaminol (Tokyo) Spec No: 585–588, 1992. 43. O Pastoris, S Savasta, P Foppa, M Catapano, M Dossena. Pyruvate dehydrogenase deﬁciency in a child responsive to thiamine treatment. Acta Paediat 85:625–628, 1996.

27 Kinetic Studies of Human Pyruvate Dehydrogenase and Its Mutants: Interactions with Thiamine Pyrophosphate Mulchand S. Patel and Lioubov G. Korotchkina School of Medicine and Biomedical Sciences, State University of New York at Buﬀalo, Buﬀalo, New York, U.S.A.

I. INTRODUCTION Pyruvate dehydrogenase (E1), the ﬁrst component of the pyruvate dehydrogenase complex (PDC), catalyzes the ﬁrst and rate-limiting step of the oxidative decarboxylation of pyruvic acid, i.e., decarboxylation of pyruvic acid and reductive acetylation of the lipoyl moieties of dihydrolipoamide acetyltransferase (E2). Lipoyl moieties are attached through the speciﬁc lysyl residues of the lipoyl domains of E2. E2 transfers acetyl groups to CoA, with the formation of acetyl-CoA, and reduced lipoyl moieties are reoxidized by dihydrolipoamide dehydrogenase (E3), with reduction of NAD+ to NADH. Besides three catalytic components, PDC has one structural component, E3binding protein (BP), that binds E3 with the central core of PDC composed of the inner domains of E2 and BP. E1 binds to the E1-binding domain of E2. Two regulatory enzymes, pyruvate dehydrogenase kinase (PDK) and phospho-pyruvate dehydrogenase phosphatase (PDP), modulate activity of PDC by phosphorylation (inactivation) and dephosphorylation (activation) of E1, respectively (1). Three serine residues are phosphorylated in the a subunit of 485

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E1: Ser264 (site 1), Ser271 (site 2), and Ser203 (site 3); phosphorylation of any one of them results in the inactivation of E1. Four isoenzymes of PDK and two isoenzymes of PDP are identiﬁed in mammals with diﬀerent speciﬁc activities and sensitivity to regulators as well as tissue-speciﬁc distribution (2–4). PDC-E1 in eukaryotes and Gram-positive bacteria is a heterotetramer composed of two a and h subunits. E1 has two active sites lying at the interface of each a and h subunit. E1 catalysis requires thiamine pyrophosphate (TPP) as a cofactor that is bound to the enzyme with participation of Mg2+. Three-dimensional structures were determined for several TPPdependent enzymes, namely, yeast transketolase (5), Lactobacillus plantarum pyruvate oxidase (6), yeast (7,8) and Zymomonas mobilis pyruvate decarboxylase (9), Pseudomonas putida benzoylformate decarboxylase (10), human and Pseudomonas putida branched-chain a-keto acid dehydrogenase (11,12), Desulfovibrio africanus pyruvate:ferredoxin oxidoreductase (13), and E. coli PDC-E1 (14), showing signiﬁcant similarity in the structures of TPP-binding regions. The common features of the active sites of these enzymes are: (1) the active sites are localized on the interface of two subunits; (2) two residues of the TPP motif (a sequence of 30–33 amino acid residues conserved in all known TPP-requiring enzymes) coordinate binding of the pyrophosphate moiety of TPP through interactions with a divalent cation (Mg2+ or Ca2+); (3) TPP is maintained in its V-conformation by several amino acid residues, among them are phenylalanine or tyrosine residues forming stacking interactions with the aminopyrimidine ring of TPP and a hydrophobic residue between two aminopyrimidine and thiazole rings; and (4) the N1V of the pyrimidine ring binds to a glutamyl residue through a hydrogen bond, which aﬀects the reactivity of 4V-amino group of TPP and activates the C2 hydrogen of the thiazolium ring of TPP. PDC-E1 structure is reported so far only for E. coli E1 (14), having a structure of a homodimer. Human PDC-E1 structure has recently been solved (14a). The structure–function relationships of E1 from human and E. coli have been studied intensively using site-directed mutagenesis (15–23). In this chapter we discuss the recent developments in the structure– function relationships of human E1, including kinetic analysis of the sitedirected mutants of E1 and regulation of E1 (and hence PDC) activity by phosphorylation–dephosphorylation. II. KINETIC ANALYSIS OF SEVERAL SUBSTITUTION MUTATIONS OBSERVED IN E1-DEFICIENT PATIENTS PDC plays an important role in glucose homeotasis; hence, deﬁciency of PDC results in congenital lactic acidosis, with severe clinical manifestations

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(24–26). The central nervous system is aﬀected the most, because glucose is its major source of energy. Symptoms vary from mild ataxia to severe mental retardation, motor function abnormalities, and, often, early death. From about 170 cases of reported PDC deﬁciency, about 130 patients had deﬁciency of E1. The other PDC components having genetic defects are E3, BP, PDP, and E2. All genetic defects of E1 were found in its a subunit and none in the h subunit. Two isoforms of E1a are present in mammalian tissues: a somatic form, encoded by the PDHA1 gene, localized on the X chromosome, and a testis-speciﬁc form, encoded by the PDHA2 gene localized on chromosome 4 (27). No defects are reported so far in the PDHA2 gene. The presence of the PDHA1 gene on the X chromosome results in a more severe manifestation in males and a high variability of severity in females because of the random X chromosome inactivation. Mutations in the PDHA1 gene are either point mutations (about onehalf) or insertions/deletions, leading to frame shifts. Mutations occur in all exons except exon 2. Missense mutations aﬀect E1 activity to diﬀerent extents. The eﬀects of several substitutions on the structure/function of E1 were investigated by recreating mutations and either studying proteins puriﬁed after overexpression in bacterial cells or testing the recovery of the PDC activity in an E1a-deﬁcient cell line after transfection of the mutant cDNAs (Table 1). Mutations of the E1-deﬁcient patients studied in the puriﬁed protein system (numbering in the mature protein) are H15, A170, M181, P188, R234, and R349 (15,17,18,28). Two of these residues, M181 and P188, reside within the TPP motif, which is a common sequence motif found in all TPP-requiring enzymes (29). It is composed of about 30 amino acid residues starting and ending with highly conserved sequences: putative –GDG– and –NN–, respectively. Aspartate and second asparagine in these sequences are involved in binding of the pyrophosphate moiety of TPP through the divalent ion (Mg2+ or Ca2+). Patients carrying mutations at M181 or P188 had severe clinical manifestations. The amount of M181V protein in the patient’s ﬁbroblasts detected by Western blotting was similar to a control subject; however, the amounts of P188L E1 proteins (both a and h proteins) were signiﬁcantly reduced, and the patient with this mutation died at the age of 2 weeks (30). Recombinant P188L E1 was very unstable (barely detectable) when overexpressed in E. coli, and hence a P188A substitution was studied instead [the stability of P188A E1 was only 5% of that observed for the wild-type (WT) E1]. Both recombinant M181V and P188A had low activity when reconstituted in PDC (Table 2) (15). Apparent Km values for TPP and pyruvate were not signiﬁcantly changed for P188A; however, the M181V substitution resulted in a 252-fold increase in the apparent Km for TPP and reduction of catalytic eﬃciency of this mutant to 0.15% of normal. WT human E1 has a lag

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Table 1 Site-Directed Mutants of Human E1

Subunit a

h

Residue

Substitution

Overexpressed in E. coli and puriﬁed

Expressed in cultured cells

H15

A Q R

+ + +

C62

A

+

H63

D N

+ +

36

H84

A D K

+ + +

36

H92

D N

+ +

36

R112

L

A170

T

+

M181

V

+

P188

A L

+

S203

A E

+ +

R234

G Q

+ +

H263

A D K L

Identiﬁed in patients

Ref. 17

+ 15

+

+

37

+

+

27, 39

+

14 14

+ 48 +

+ +

+ + +

16, 39 36, 39

+

S264

A E Q D

+ + + +

48

S271

A E Q

+ + +

48

R273

C

R349

H

+

E59

D Q A

+ + +

22

W135

L

+

15

+

+

38

+

14

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Table 2 Kinetic Parameters of E1 Mutants

E1

Mutation

WT

None

Activity (%) 100

Increase in Km for TPP (fold) 1

Increase in Km for Pyr (fold) 1

Identiﬁed in patients M181V P188A R349H R234G R234Q H15R

38.5 22.3 1.2 126.8 97.0 6.0

252 0.9 ND 6.8 1.4 2.8

2.9 1.3 ND 0.9 0.8 1.1

Substitutions H15Q H15A W135L C62A

30.9 86.8 11.2 7.9

6.6 10.4 5.9 5.9

1.2 1.0 1.5 2.1

Mutant E1s were ﬁrst reconstituted in PDC by preincubation with recombinant human E2-BP and recombinant human E3, and the activity was measured using the PDC assay by formation of NADH. One hundred percent of activity corresponds to 28 F 2 units/mg of E1. Km for TPP is 80 nM and Km for pyruvate is 52 AM when determined in PDC assay. ND, not determined.

phase in the progress curve of the PDC reaction, depending on the TPP concentration, reﬂecting activation of E1 during catalysis (15). Lag phase was not seen for the WT E1 at TPP concentrations higher than 1 lM, but it was still present and independent on TPP concentration for M181V at more than 500 lM TPP and for P188A at more than 1 lM TPP (Fig. 1). This ﬁnding indicates that these mutations have changed the conformation of the TPPbinding site in such a way that complete activation of E1 is reached only during catalysis and not with saturation with TPP. Thiamine thiothiazolone pyrophosphate (TTTPP) was shown to be a slow and tight-binding inhibitor of human E1 (15). Preincubation with TTTPP inactivated E1 (Fig. 2A); however, TPP and TPP plus pyruvate protected E1 from inactivation and could restore E1 activity when added after preincubation with TTTPP (Fig. 2B). Stoichiometry of TTTPP binding was 2.3 TTTPP/E1 for the WT E1, 1.1 TTTPP/E1 for P188A E1, and 4.3 TTTPP/E1 for M181V. The aﬃnity of M181V for TTTPP was decreased, similar to its reduction in aﬃnity to TPP. P188 is positioned on the turn between two structural elements (a-helix and h-sheet) of the TPP motif. Its integrity could determine the conformation

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Figure 1 Dependence of the lag-phase duration on TPP concentration. Lag phase was determined from the progress curves of the PDC reaction at the indicated diﬀerent TPP concentrations for WT (A, . ), P188A (A, o), and M181V (B, E). Reactions were started by the addition of TPP. Lag phase was calculated using the equation P(t) = vsst H (vss v0)(1exp(t/H )), where P(t) is the product concentration, vss is the steady-state reaction rate, H is the lag-phase duration, v0 is the initial rate, and t is the time of observation.

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Figure 2 Inactivation of WT and mutant E1s by thiamine thiothiazolone pyrophosphate (TTTPP). (A) WT (.), P188A (n), and M181V (D) were preincubated with 2 mM MgCl2 and diﬀerent amounts of TTTPP for 15 min. Aliquots were taken and activity was determined in the PDC assay. (B) Eﬀect of TPP and pyruvate on TTTPP inactivation: 1, before treatment; 2, after inhibition by 16.7 lM TTTPP; 3, inhibition by 16.7 lM ThTTPP in the presence of 10 mM TPP; 4, inhibition in the presence of 10 mM pyruvate; 5, inhibition in the presence of TPP and pyruvate; 6, reactivation by 30min incubation with 10 mM TPP after 30-min inhibition by TTTPP; and 7, reactivation by 10 mM TPP plus 10 mM pyruvate after inhibition by TTTPP. White bars, untreated E1; grey bars, inhibition of E1 with TTTPP in the absence or presence of TPP and pyruvate; black bars, inhibition of E1 and following reactivation by TPP and pyruvate.

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of the whole TPP motif. Mutation at this position could increase the instability of the conformation of the TPP-binding site, resulting in an unstable and less active E1. M181 could be involved in the hydrophobic interaction with the neighboring residues, and its mutation would aﬀect the position of the TPP motif and its two residues, Asp167 and Asn196, which coordinate the binding of the pyrophosphate moiety of TPP through interactions with Mg2+. This would explain the observed reduction in the aﬃnity for TPP. An R349H mutation was found in six unrelated patients. Activity of the recombinant R349H mutant E1 determined after its reconstitution in PDC was drastically reduced to the level of about 1.2% as compared to WT, indicating an essential role of this residue in E1 function (Table 2) (15). R349 is localized at the C-terminus of the E1a subunit and could be involved in the interaction between the a and h subunits. Perturbation of this interaction could aﬀect the active site of E1, localized at the interface between the two subunits. R234 is a hot spot of mutations. An R234G substitution was found in 14 patients and four of their mothers (31–33). Furthermore, an R234Q mutation was reported in three patients. Interestingly, the recombinant R234G mutant E1 was as active as the WT E1 but had an apparent Km value for TPP approximately sevenfold higher than that for the WT E1. Decreased aﬃnity for TPP could explain PDC deﬁciency in aﬀected patients, and thiamine treatment was shown to be eﬀective for one patient with the R234G mutation (34). Kinetic parameters of E1 with another substitution at the same position, R234Q, were not signiﬁcantly diﬀerent from those of the WT E1 (Table 2) (17). The diﬀerent speciﬁcities of PDK isoenzymes for the three phosphorylation sites provided approach for the investigation of the eﬀect of the R234Q mutation on phosphorylation/dephosphorylation of site 1, site 2, and site 3 individually. Only PDK1 phosphorylates site 3 (35). Phosphorylation of sites 1 and 2 was carried out in the presence of PDK2, followed by dephosphorylation catalyzed by PDP1. When phosphorylation of site 3 was tested, E1 was prephosphorylated on sites 1 and 2 with unlabeled ATP (Fig. 3). The modiﬁcation of site 3 was then performed by PDK1 in the presence of [g-32P]ATP, followed by dephosphorylation by PDP1. Figure 3 shows that the rates of phosphorylation of sites 1 plus 2 and site 3 as well as the rates of dephosphorylation of sites 1 plus 2 were similar among the WT, R234G, and R234Q E1s. However, the rate of dephosphorylation of site 3 of R234Q was only about 50% of WT (Fig. 3). Dephosphorylation of all three sites is necessary for E1 reactivation, and hence a decrease in the rate of dephosphorylation of one of the phosphorylation sites would result in reduction in the PDC activity in these patients. Three strategies are used for the therapy of E1 deﬁciency: (1) high-fat diets with limited amounts of carbohydrates; (2) administration of dichloro-

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Figure 3 Eﬀect of glycine or glutamine substitution of R234 on the rates of phosphorylation and dephosphorylation of E1s. (A) Apparent rate constants of phosphorylation of WT, R234G, and R234Q sites 1 plus 2 (open columns) and site 3 (hatched columns). Sites 1 and 2 were phosphorylated simultaneously by PDK2 in the presence of [g-32P]ATP. For site 3, phosphorylation sites 1 and 2 were prephosphorylated by PDK2 in the presence of unlabeled ATP, and site 3 was then phosphorylated by PDK1 in the presence of [g-32P]ATP (as depicted in the ﬂowchart shown on the right-hand panel). (B) Apparent rate constants of reactivation of WT, R234G, and R234Q phosphorylated at sites 1 and 2 by PDP1 as determined by the restoration of E1 activity in PDC assay (open columns) and dephosphorylation of site 3 determined by the decrease in residual radioactivity of E1 (hatched columns).

acetate to inhibit PDK and hence decrease the level of E1 in the phosphorylated (inactive) form; and (3) high doses of thiamine (25). Patients with several mutations (R234G, C72F, V42A, and H15R) were reported to be responsive to thiamine treatment (34,36). To study the role of H15, several substitutions were created: H15R, H15Q, and H15A (18). Recreation of the patient’s mutation, H15R, resulted in a very low activity of about 6% as compared to the WT, with a moderate increase (2.8-fold) in the apparent Km for TPP (Table 2). The H15A mutant E1 had near-normal activity but a very large increase (10-fold) in the apparent Km for TPP. Apparently, an increase in the size of the side chain of the residue substituting for the histidine at this site

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caused a higher level of reduction in activity, indicating a steric constraint for H15. The dependence of the Km for TPP on the substitutions at H15 is the opposite of the speciﬁc activity dependence. Alanine substitution caused the highest increase in the Km for TPP, while the change in the Km for TPP for the H15R mutant E1 was only moderate. It is possible that the positive charge of H15 is important for the TPP binding and that its complete elimination results in a decreased TPP binding. Interestingly, H15 is localized near the N-terminus of the E1a subunit, which is not close to the active site2. However, substitutions of H15 resulted in an increase in the apparent Km for TPP, which correlated with an improvement in the patient’s clinical condition when thiamine was administrated (36). H15 may play a structural role, and its substitution may cause disturbance in the TPP-binding site. Recombinant A170T mutant E1 had about 25% of control activity and the apparent Km for pyruvate increased by approximately 10-fold (28). The apparent Km for TPP was increased only about two fold. This mutation may have a possible eﬀect on the binding of pyruvate. Another approach to study the roles of the mutations identiﬁed in the E1-deﬁcient patients as well as other tested mutations is to express recombinant mutant E1a in a human ﬁbroblast cell line with no endogenous E1a mRNA and protein. Impairment of the E1 function is determined by the extent to which the mutant E1a could restore PDC activity as compared to the restoration by WT E1. Using this system it was demonstrated that mutants H63D, H63N, H92D, H92N, R112L, H263A, H263D, H263K, and R273C did not have E1 activity (37–39). Other mutations caused less severe impairment of the E1 function: Reconstitution of mutant A170T resulted in 17% of PDC activity, expression of H84A, H84D, H84K gave about 33% of PDC activity, and the R234G mutant E1 produced PDC with 39% activity (37,40). All mutants were expressed at the same level as WT E1a except H92 (both substitutions), indicating a possible eﬀect of the H92 substitution on: (1) the import of E1a into mitochondria, (2) the stability of mutant E1a in the cell, or (3) an inability of mutant E1a to interact with E1h during PDC assembly that will lead to E1a degradation (37). The results for A170T mutant E1 correlated with the E1 activity detected in the ﬁbroblasts of the corresponding patient (17%), while mutations R273C and R234G showed a broad range of activity determined in the patient’s tissues: 10–50% and 16–168%, respectively, as compared to normal (26). III. MUTAGENESIS STUDY OF SEVERAL ESSENTIAL AMINO ACID RESIDUES IN E1 Several amino acid residues were found to be essential for mammalian E1 activity, based on chemical modiﬁcation of speciﬁc cysteine, tryptophan,

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lysine, arginine, and histidine residues (41–45). Three of them were identiﬁed by chemical modiﬁcation and diﬀerential peptide mapping: C62 in the E1a subunit and W135 and R239 in the E1h subunit (41,43,45). aC62 and hW135 were studied further by employing site-directed mutagenesis (16). The aC62A and hW135L mutations resulted in a signiﬁcantly reduced activity of E1, determined in both PDC and 2,6-dichlorophenolindophenol (DCPIP) assays (Table 2). Kinetic analysis showed that the apparent Km value for pyruvate did not change signiﬁcantly, whereas the apparent Km value for TPP increased by about six fold for both mutants. This resulted in the low catalytic eﬃciency for both aC62A (1.2%) and hW135L (1.9%). Both mutant proteins were less thermostable than the WT E1. The half-lives of aC62A and hW135L at 37jC were 24% and 15%, respectively, as compared to that of the WT E1. Spectral studies (CD) of the E1 mutant proteins revealed signiﬁcant diﬀerences in the conformations of aC62A and hW135L as compared to the WT E1 as well as impairment in TPP binding. Diﬀerences in TPP binding with the mutant E1s were also reﬂected in the presence of the lag phase in the progress curve of the PDC reaction, even at high concentrations of TPP, at which no lag phase was detected in the WT E1 (15). hW135 is most probably not localized in the vicinity of the E1 active site but rather plays a structural role. hW135L may have changed conformation, which is causing less eﬃcient binding of TPP. This mutation results in a less stable protein that may dissociate into dimers or monomers during prolonged exposure to higher temperature. aC62 may be localized close to the active site of E1. Human E1 was shown to be inhibited by the substrate analogs, bromopyruvate and ﬂuoropyruvate, undergoing reaction in the presence of TPP in the E1 active site. The adduct formed during this reaction could be trapped on amino acid residue(s) of the active site. The substrate of the E1 reaction, pyruvate, causes E1 inactivation in the presence of TPP and the absence of the electron acceptors. E1 could be reactivated after pyruvate-induced inactivation by the reducing reagents (dithiothreitol, cysteine). The aC62A mutant E1 neither was inhibited by the site-directed inhibitors, bromopyruvate and ﬂuoropyruvate, nor underwent the substrate-induced inactivation (16). Both phenomena are suggested to involve a cysteinyl residue as a participant (42,46). It is possible that aC62 is one of the residues involved in both types of inhibition. That places aC62 close to the pyruvate-binding site and explains the reduced aﬃnity for TPP. Another catalytic residue studied by means of site-directed mutagenesis is hE59. The presence of a glutamyl residue near the N1V position of the aminopyrimidine ring of TPP is the common feature of the TPP-dependent enzymes. This residue is necessary for the activation of TPP during catalysis. Replacement of hE59 with alanine, glutamine, or aspartate resulted in a

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signiﬁcantly lower aﬃnity for TPP and the reduction of E1 speciﬁc activity to 1, 5, and 13%, respectively (23). IV. REGULATION OF E1 ACTIVITY BY PHOSPHORYLATION Activity of PDC is regulated in response to changes in the concentrations of metabolites and hormones by the modulation in the level of phosphorylation of its ﬁrst component, E1. Phosphorylation of E1 resulting in inactivation is catalyzed by a speciﬁc kinase, PDK, and dephosphorylation (activation) is catalyzed by a speciﬁc phosphatase, PDP. Regulation of the activities of PDK, and PDP determines the level of phosphorylation of E1 and hence the level of active PDC (1,3,4). The long-term regulation of PDC activity during pathophysiological conditions (disease, nutritional state) is also achieved by the modulation of the amounts of PDC components at the transcriptional level. Up-regulation of PDK and down-regulation of PDP during diabetes and starvation play an important role in PDC regulation (4,47). Regulation of PDC by phosphorylation–dephosphorylation is found so far in humans, rodents, plants, nematodes, and the fruit ﬂy (for a review, see Ref. 2). Three speciﬁc serine residues are phosphorylated by PDKs in mammalian tissues and are identiﬁed as site 1 (Ser264), site 2 (Ser271), and site 3 (Ser203), based on their rates of phosphorylation in mammalian E1. Isoform II of Ascaris suum E1 and fruit ﬂy E1 also have three sites of phosphorylation, whereas only two phosphorylation sites are present in isoform I of Ascaris suum E1 and Caenorhabditis elegans E1 and only one site is present in plant E1s. Phosphorylation site 1 is present in the sequence of yeast E1, and its phosphorylation by bovine PDK results in inactivation; however, no PDK was detected in yeast. Site-directed mutagenesis was employed to study site-speciﬁc phosphorylation of human E1 (48). It was found that each of the three sites was phosphorylated either individually or in combination with the other sites. Phosphorylation of each site resulted in E1 inactivation. The mechanism of inactivation was studied further using kinetic analysis of the mutant E1s, with several substitutions of the serine residues of the phosphorylation sites (49). Substitution of site 1 with either glutamate or aspartate caused complete inactivation of E1 when PDC activity was determined (Fig. 4A, Table 3), similar to phosphorylation of site 1. The glutamine-substituted mutant of site 1 had about 3% activity in PDC assay, indicating that not only the introduction of the negative charge but also an increase in the size of the amino acid residue in position 264 reduces E1 activity. Even alanine substitution at site 1 resulted in about a 40% reduction in PDC activity. Interestingly, mutants of site 1 only and E1 phosphorylated on site 1 had signiﬁcant levels of activity in the decarboxylation assay (measuring only the ﬁrst

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Figure 4 Activities of the WT, glutamate-substituted E1 mutants of sites 1 (S1E), 2 (S2E), and 3 (S3E) and WT E1 phosphorylated at site 1, 2, or 3. Activities were measured in PDC assay (black bars) and decarboxylation assay (open bars). (A) Activities for WT (unphosphorylated as a control), S1E (site 1), S2E (site 2), and S3E (site 3). (B) Activities of WT (unphosphorylated), site 1 (E1-S2A-S3A mutant phosphorylated at site 1 by PDK2), site 2 (E1-S1A-S3A phosphorylated at site 2 by PDK2), and site 3 (E1-S1A-S2A phosphorylated at site 3 by PDK1).

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Table 3 Kinetic Parameters of E1 Phosphorylation Mutants Increase in Km for Pyr (fold)

Site

Mutant

Activity (%)

Increase in Km for TPP (fold)

1

S264A S264E S264Q

57.9 0 3.3

12.5 ND (1, DCPIP) ND (1.7, DCPIP)

2.3 ND (4.5, DCPIP) ND (2.6, DCPIP)

2

S271A S271E S271Q

62.9 45.4 51.3

2.4 9.8 3.3

0.8 1.5 1.1

3

S203A S203E

59.0 48.2

1.1 27

1.0 2.8

Unless otherwise indicated, the activity of mutant E1s was measured as PDC after reconstituting in the complex. ND, not determined. DCPIP, assayed using 2,6-dichlorophenolindophenol.

partial reaction of E1, i.e., decarboxylation of [1-14C]pyruvate to 14CO2) (Fig. 4) or DCPIP assay (Fig. 5A). This ﬁnding suggested a larger impact on the second partial reaction catalyzed by E1, i.e., reductive acetylation of the lipoyl moieties of E2 as a result of site 1 phosphorylation. Apparent Km values for pyruvate determined in DCPIP assay (required because of the absence or very low activities of site 1 mutants in PDC assay) increased three- to ﬁvefold for alanine-, aspartate-, and glutamate-substituted E1 mutants of site 1. Ser264 may participate in catalysis through interactions with the substrates and intermediates. Phosphorylation of Ser264 may distort interactions with pyruvate and even more with the lipoyl domain of E2 because of the volume and negative charge of the phosphoryl group. Phosphorylation changes the substrate recognition site because replacing serine264 with alanine, glutamine, or glutamate leads to an increase in the mutant E1 activity with a-ketobutyrate as a substrate, reaching 500% with glutamate-substituted mutant E1 (Fig. 5B). Phosphorylation of site 2 or 3 caused a drastic reduction in activity measured in PDC assays and only moderate reduction (about 40%) in activity measured in the decarboxylation reaction (Fig. 4B). Glutamate substitution of site 2 or site 3 resulted in 40–55% reduction in activity measured in both the assays. Changes in the local conformation near sites 2 and 3 during protein folding could explain partial compensation of the activity loss during phosphorylation. Apparent Km values for pyruvate did not change signiﬁ-

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Figure 5 Activities of WT and phosphorylation site mutant E1s with 5 mM pyruvate (A) and 5 mM a-ketobutyrate (B) as the substrate, determined with 2,6dichlorophenolindophenol (DCPIP assay). 1A, 1E, 1Q: E1 mutants with site 1 substituted with alanine, glutamate, and glutamine, respectively; 2A, 2E, 2Q: E1 mutants with site 2 substituted with alanine, glutamate, and glutamine, respectively; 3A, 3E: E1 mutants with site 3 substituted with alanine and glutamate, respectively. One hundred percent of WT activities corresponded to 242 mU/mg with pyruvate and 34 mU/mg with a-ketobutyrate. Results are means F S.D. of four to ﬁve independent determinations.

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cantly for the mutants of sites 2 and 3. However, apparent Km values for TPP were approximately 10-fold and 27-fold higher for the glutamate-substituted site 2 and site 3 E1 mutants, respectively, as compared to the WT E1. TPP protected sites 1, 2, and 3 from phosphorylation by PDK1. Protection was higher for site 3 than for site 2, and higher for site 2 than for site 1. Site 3 could be localized close to the TPP-binding region, and its phosphorylation may aﬀect TPP binding. Another eﬀect of phosphorylation on E1 activity in PDC could be that the negative charge introduced at site 2 or 3 may prevent binding of the lipoyl domain. The region of lipoyl domain determining speciﬁcity of binding to E1 includes several acidic residues (3). Several isoforms of PDK are identiﬁed, four in mammalian PDC (PDK1, PDK2, PDK3, and PDK4), two in plants, and one in nematodes and the fruit ﬂy (50–55). Isoenzymes of PDK are diﬀerent in speciﬁc activities and sensitivity to diﬀerent eﬀectors (acetyl-CoA, NADH, pyruvate, ADP) (3). Tissue-speciﬁc distribution of PDK isoenzymes allows selective modulation of PDC activity, depending on the role of a speciﬁc tissue in the maintenance of glucose homeostasis. PDK1 mRNA was detected mostly in heart and less in skeletal muscle, liver, and pancreas. PDK2 is expressed in many tissues, with low levels in spleen and lung. PDK3 mRNA is high in testis and less in lung, brain, and kidney. PDK4 mRNA is present in skeletal muscle and heart, with low levels in lung, liver, and kidney (50,51,56). During the long-term regulation of PDC activity, the amounts of two isoenzymes, PDK2, and PDK4, were found to increase (4,57). The levels of PDK4 increase during starvation in liver, kidney, lactating mammary gland, heart, and muscle, and during diabetes and hyperthyroidism in heart and skeletal muscle. The levels of PDK2 are higher during starvation in liver, kidney, lactating mammary gland, fast-twitch muscle, slightly elevated during diabetes in skeletal muscle, and are increased during hyperthyroidism in fast-twitch muscle. The elevated levels of fatty acids in certain physiological conditions are proposed to increase expression of PDK4 through activation of PPARa in liver cells (58). However, overexpression of PDK4 in muscle and heart did not depend on signaling through PPARa, indicating that the mechanisms of PDK4 upregulation are diﬀerent in diﬀerent tissues (59). Up-regulation of PDK expression could result in phosphorylation at the multiple sites of E1, which would lead to a longer duration of inactivation of PDC. PDK isoenzymes have diﬀerent activities toward the three phosphorylation sites of E1 (Fig. 6) (35). PDK1 can phosphorylate all three sites in E1, whereas PDK2, PDK3, and PDK4 modify only sites 1 and 2. PDK2 had the highest activity toward site 1, and PDK3 phosphorylated site 2 faster than other PDKs (Fig. 6). PDK3 had higher activity toward site 2 than site 1. PDKs are bound to the lipoyl domains of E2 or BP in PDC: PDK1 binds to

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Figure 6 Activities of PDK isoenzymes determined using reconstituted PDC (in the presence of E2-BP) or free E1 (in the absence of E2-BP). E1 mutants used as substrates for PDKs: E1-S2A-S3A for site 1 phosphorylation, E1-S1A-S3A for site 2 phosphorylation, and E1-S1A-S2A for site 3 phosphorylation. E1s were reconstituted with E2-BP, E3, and PDK into PDC and incubated for 5 min with NAD+/ NADH =794/4 lM before the reaction was started by the addition of [g-32P]ATP. Activity is expressed as nmoles of 32P incorporated per milligram of PDK per minute. Results are means F S.D. of three to six independent determinations.

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L1 and L2 in E2; PDK2 and PDK3 prefer L2 but can also bind L1 less eﬃciently; and PDK4 is bound to L3 in BP and less to L1 (3). Binding of PDK to the lipoyl domain of E2 activates it. The activation may be achieved by the conformational change(s) of PDK when bound to the lipoyl domain and colocalization of PDK and its substrate E1 (60). The degree of activation is diﬀerent for diﬀerent isoenzymes of PDK. The maximum activation is observed for PDK3. Activities of PDKs are regulated by reduction of the lipoyl moieties of E2. PDK activity is stimulated with the reduction and reduction/acetylation of the lipoyl groups of E2. The stimulation is explained by a conformational change in PDK upon reduction or acetylation of the lipoyl moiety of the lipoyl domain. The extent of stimulation depends on the PDK isoenzyme (highest for PDK2) and not on the involvement of the phosphorylation site (3,35). Interestingly, PDKs can phosphorylate free E1 as a substrate. All four PDKs phosphorylated site 1 and site 2, and PDK1 also modiﬁed site 3 using free E1. Activity of PDK4 was higher toward site 2 of free E1 than of the other isoenzymes (35). This could play a role during certain pathophysiological conditions when the amount of PDK4 is increased. V. CONCLUSIONS Kinetic analysis of E1 mutants, either naturally occurring or created based on predictions, provided an approach to investigate the role of a particular amino acid residue in the structure and function of E1. Several mutations were found to be essential for E1 activity with altered kinetic parameters. Only one of the mutants studied so far, R234Q, appeared to have an impact not on the function of E1 but on its regulation, i.e., decreasing the rate of dephosphorylation of site 3. Most of the mutations aﬀected the Km for TPP, although most of the residues were not localized in the active site and were not directly involved in TPP binding. These ﬁndings indicate that: (1) the TPPbinding site is the ‘‘heart’’ of the E1 protein, sensing even remote changes in the structure, and (2) not only residues forming the active site and directly involved in the interactions with the coenzyme and the substrates are essential, but a whole network of residues is involved in interacting with each other to provide a conformation for high catalytic eﬃciency and sensitivity to regulation. The complex mechanism of regulation of PDC activity by phosphorylation–dephosphorylation of the three phosphorylation sites of E1 by a family of tissue-speciﬁc PDKs and PDPs with participation of E2, binding E1 as well as PDK and PDP and modulating their activities by the redox state, provides the immediate as well as the long-term responses to the metabolic state of the cell.

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ACKNOWLEDGMENTS This work was supported by U.S. Public Health Service Grant DK20478. We are grateful to Dr. Murray Ettinger of the Department of Biochemistry at SUNY Buﬀalo for his critical reading of the manuscript and helpful discussions.

ABBREVIATIONS PDC E1 E2 E3 BP PDK PDP TPP WT ThTTPP E1-S1E, E1-S2E E1-S3E E1-S2A-S3A E1-S1A-S3A E1-S1A-S2A DCPIP PPARa

pyruvate dehydrogenase complex; pyruvate dehydrogenase; dihydrolipoamide acetyltransferase dihydrolipoamide dehydrogenase E3-binding protein pyruvate dehydrogenase kinase phospho-pyruvate dehydrogenase phosphatase thiamine pyrophosphate wild-type thiamine thiothiazolone pyrophosphate E1 with phosphorylation site 1 serine substituted by glutamate E1 with phosphorylation site 2 serine substituted by glutamate E1 with phosphorylation site 3 serine substituted by glutamate E1 with serines at sites 2 and 3 substituted by alanine E1 with serines at sites 1 and 3 substituted by alanine E1 with serines at sites 1 and 2 substituted by alanine 2,6-dichlorophenolindophenol peroxisome proliferator-activated receptor a

REFERENCES 1. 2.

3.

MS Patel, TE Roche. Molecular biology and biochemistry of pyruvate dehydrogenase complexes. FASEB J 4:3224–3233, 1990. MS Patel, LG Korotchkina. Regulation of mammalian pyruvate dehydrogenase complex by phosphorylation: complexity of multiple phosphorylation sites and kinases. Exp Mol Med 33:191–197, 2001. TE Roche, JC Baker, X Yan, Y Hiromasa, X Gong, T Peng, J Dong, A Turkan, SA Kasten. Distinct regulatory properties of pyruvate dehydrogenase kinase and phosphatase isoforms. Progr Nucl Acid Res Molec Biol 70:33–75, 2001.

504

Patel and Korotchkina

4.

RA Harris, B Huang, P Wu. Control of pyruvate dehydrogenase kinase gene expression. Adv Enzyme Regul 41:269–288, 2001. 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. 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. F Dyda, W Furey, S Swaminathan, M Sax, B Farrenkopf, F Jordan. Catalytic centers in the thiamin diphosphate–dependent enzyme pyruvate decarboxylase at 2.4-A˚ resolution. Biochemistry 32:6165–6170, 1993. P Arjunan, T Umland, F Dyda, S Swaminathan, W Furey, M Sax, B Farrenkorf, Y Gao, D Zhang, F Jordan. Crystal structure of the thiamin diphosphate–dependent enzyme pyruvate decarboxylase from the yeast Saccaromyces cerevisiae at 2.3-A˚ resolution. J Mol Biol 256:590–600, 1996. D Dobritzsch, S Konig, 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. 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. A Aevarsson, JL Chuang, RM Wynn, S Turley, DT Chuang, WGJ Hol. Crystal structure of human branched-chain a-ketoacid dehydrogenase and the molecular basis of multienzyme complex deﬁciency in marple syrup urine disease. Structure 8:277–291, 2000. A Aevarsson, K Seger, S Turley, JR Sokatch, WGJ Hol. Crystal structure of 2-oxoisovalerate and dehydrogenase and the architecture of 2-oxo acid dehydrogenase multienzyme complexes. Nat Struct Biol 6:785–792, 1999. E Chabriere, MH Charon, A Volbeda, L Pieulle, EC Hatchikian, JC FontecillaCamps. Crystal structures of the key anaerobic enzyme pyruvate:ferredoxin oxidoreductase, free and in complex with pyruvate. Nat Struct Biol 6:182–190, 1999. 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. EM Ciszak, LG Korotchkina, PM Dominiak, S Sidhu, MS Patel. Structural basis for ﬂip-ﬂop action of thiamin pyrophosphate-dependent enzymes revealed by human pyruvate dehydrogenase. J Biol Chem 278:21240–21246, 2003. A Tripatara, LG Korotchkina, MS Patel. Characterization of point mutations in patients with pyruvate dehydrogenase deﬁciency: role of methionine-181, proline-188, and arginine-349 in the alpha subunit. Arch Biochem Biophys 367:39–42, 1999. LG Korotchkina, MS Ali, MS Patel. Involvement of a-cysteine-62 and b-tryptophan-135 in human pyruvate dehydrogenase catalysis. Arch Biochem Biophys 369:277–287, 1999.

5. 6.

7.

8.

9.

10.

11.

12.

13.

14.

14a.

15.

16.

Kinetic Studies of Human E1

505

17. SJ Jacobia, LG Korotchkina, MS Patel. Diﬀerential eﬀects of two mutations at arginine-234 in the alpha subunit of human pyruvate dehydrogenase. Arch Biochem Biophys 395:121–128, 2001. 18. SJ Jacobia, LG Korotchkina, MS Patel. Characterization of a missense mutation at histidine-44 in a pyruvate dehydrogenase–deﬁcient patient. Biochim Biophys Acta 1586:32–42, 2002. 19. GC Russell, RS Machado, JR Guest. Overproduction of the pyruvate dehydrogenase multienzyme complex of Escherichia coli and site-directed substitutions in the E1p and E2p subunits. Biochem J 287:611–619, 1992. 20. J Yi, N Nemeria, A McNally, F Jordan, RS Machado, JR Guest. Eﬀect of substitutions in the thiamin diphosphate–magnesium fold on the activation of the pyruvate dehydrogenase complex from Escherichia coli by cofactors and substrate. J Biol Chem 271:33192–33200, 1996. 21. N Nemeria, A Volkov, A Brown, J Yi, L Zipper, JR Guest, F Jordan. Systematic study of the six cysteines of the E1 subunit of the pyruvate dehydrogenase multienzyme complex from Escherichia coli: none is essential for activity. Biochemistry 37:911–922, 1998. 22. N Nemeria, Y Yan, Z Zhang, AM Brown, P Arjunan, W Furey, JR Guest, F Jordan. Inhibition of the Escherichia coli pyruvate dehydrogenase complex E1 subunit and its tyrosine 177 variants by thiamin 2-thiazolone and thiamin 2-thiothiazolone diphosphates. Evidence for reversible tight-binding inhibition. J Biol Chem 276:45969–45978, 2001. 23. R Fang, PF Nixon, RG Duggleby. Identiﬁcation of the catalytic glutamate in the E1 component of human pyruvate dehydrogenase. FEBS Lett 437:273– 277, 1998. 24. MS Patel, RA Harris. Mammalian alpha-keto acid dehydrogenase complexes: gene regulation and genetic defects. FASEB J 9:1164–1172, 1995. 25. DS Kerr, ID Wexler, A Tripatara, MS Patel. Human defects of the pyruvate dehydrogenase complex. In: MS Patel, TE Roche, eds. Alpha-Keto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996, pp 249–269. 26. W Lissens, L De Meirleir, S Seneca, I Liebaers, GK Brown, RM Brown, M Naito, E Naito, Y Kuroda, DS Kerr, ID Wexler, MS Patel, BH Robinson, A Seyda. Mutations in the X-linked pyruvate dehydrogenase (E1) alpha subunit gene (PDHA1) in patients with a pyruvate dehydrogenase complex deﬁciency. Hum Mutat 15:209–219, 2000. 27. HHM Dahl, J Fitzgerald, R Iannelo. The sperm-speciﬁc pyruvate dehydrogenase E1a genes. In: MS Patel, TE Roche, eds. Alpha-Keto Acid Dehydrogenase Complexes. Basel: Birkhauser Verlag, 1996, pp 213–226. 28. YG Wu, SL Widjaja, CY Huang, W Li, PF Nixon, RG Duggleby. Congenital lactic acidosis: evaluation of the properties of the a199t natural variant of human pyruvate dehydrogenase e1 alpha by in vitro mutation. Mol Genet Metab 72:269–272, 2001. 29. CF Hawkins, A Borges, RN Perham. A common structural motif in thiamin pyrophosphate–binding enzymes. FEBS Lett 255:77–82, 1989. 30. SG Hemalatha, DS Kerr, ID Wexler, MM Lusk, M Kaung, Y Du, M Kolli, RL Schelper, MS Patel. Pyruvate dehydrogenase complex deﬁciency due to a

506

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

Patel and Korotchkina point mutation (P188L) within the thiamine pyrophosphate–binding loop of the E1 alpha subunit. Hum Mol Genet 4:315–318, 1995. ID Wexler, SG Hemalatha, TC Liu, SA Berry, DS Kerr, MS Patel. A mutation in the E1 alpha subunit of pyruvate dehydrogenase associated with variable expression of pyruvate dehydrogenase complex deﬁciency. Pediatr Res 32:169– 174, 1992. K Chun, N MacKay, R Petrova-Benedict, A Federico, A Fois, DE Cole, E Robertson, BH Robinson. Mutations in the X-linked E1 alpha subunit of pyruvate dehydrogenase: exon skipping, insertion of duplicate sequence, and missense mutations leading to the deﬁciency of the pyruvate dehydrogenase complex. Am J Hum Genet 56:558–569, 1995. H Awata, F Endo, A Tanoue, A Kitano, I Matsuda. Characterization of a point mutation in the pyruvate dehydrogenase E1 alpha gene from two boys with primary lactic acidemia. J Inherit Metab Dis 17:189–195, 1994. E Naito, M Ito, I Yokota, T Saijo, Y Ogawa, Y Kuroda. Diagnosis and molecular analysis of three male patients with thiamine-responsive pyruvate dehydrogenase complex deﬁciency. J Neurol Sci 201:33, 2002. LG Korotchkina, MS Patel. Site speciﬁcity of four pyruvate dehydrogenase kinase isoenzymes toward the three phosphorylation sites of human pyruvate dehydrogenase. J Biol Chem 276:37223–37229, 2001. E Naito, M Ito, E Takeda, I Yokota, S Yoshijima, Y Kuroda. Molecular analysis of abnormal pyruvate dehydrogenase in a patient with thiamineresponsive congenital lactic acidemia. Pediatr Res 36:340–346, 1994. A Seyda, BH Robinson. Functional expression of four PDH-E(1)alpha recombinant histidine mutants in a human ﬁbroblast cell line with zero endogenous PDH complex activity. Biochem Biophys Res Commun 270:1068– 1073, 2000. A Seyda, K Chun, S Packman, BH Robinson. A case of PDH-E1 alpha mosaicism in a male patient with severe metabolic lactic acidosis. J Inherit Metab Dis 24:551–559, 2001. LJ Otero, RM Brown, GK Brown. Arginine 302 mutations in the pyruvate dehydrogenase E1alpha subunit gene: identiﬁcation of further patients and in vitro demonstration of pathogenicity. Hum Mutat 12:114–121, 1998. K Chun, BH Robinson. Expression of normal and mutant pyruvate dehydrogenase complex E1 alpha cDNAs in cultured human lymphoblasts. Arch Biochem Biophys 349:246–250, 1998. MS Ali, TE Roche, MS Patel. Identiﬁcation of the essential cysteine residue in the active site of bovine pyruvate dehydrogenase. J Biol Chem 268:22353– 22356, 1993. LS Khailova, LG Korochkina, SE Severin. Organization and functioning of muscle pyruvate dehydrogenase active centers. Ann NY Acad Sci 573:36–54, 1989. MS Ali, BC Shenoy, D Eswaran, LA Andersson, TE Roche, MS Patel. Identiﬁcation of the tryptophan residue in the thiamin pyrophosphate–binding site of mammalian pyruvate dehydrogenase. J Biol Chem 270:4570–4574, 1995.

Kinetic Studies of Human E1

507

44. LR Stepp, LJ Reed. Active-site modiﬁcation of mammalian pyruvate dehydrogenase by pyridoxal 5V-phosphate. Biochemistry 24:7187–7191, 1985. 45. D Eswaran, MS Ali, BC Shenoy, LG Korotchkina, TE Roche, MS Patel. Arginine-239 in the beta subunit is at or near the active site of bovine pyruvate dehydrogenase. Biochim Biophys Acta 1252:203–208, 1995. 46. DS Flournoy, PA Frey. Inactivation of the pyruvate dehydrogenase complex of Escherichia coli by ﬂuoropyruvate. Biochemistry 28:9594–9602, 1989. 47. B Huang, KM Popov, RA Harris. Expression of pyruvate dehydrogenase phosphatase 2 is decreased in diabetes and starvation. Diabetes 51:1288, 2002. 48. LG Korotchkina, MS Patel. Mutagenesis studies of the phosphorylation sites of recombinant human pyruvate dehydrogenase. Site-speciﬁc regulation. J Biol Chem 270:14297–14304, 1995. 49. LG Korotchkina, MS Patel. Probing the mechanism of inactivation of human pyruvate dehydrogenase by phosphorylation of three sites. J Biol Chem 276: 5731–5738, 2001. 50. R Gudi, MM Bowker-Kinley, NY Kedishvili, Y Zhao, KM Popov. Diversity of the pyruvate dehydrogenase kinase gene family in humans. J Biol Chem 270: 28989–28994, 1995. 51. J Rowles, SW Schrer, T Xi, M Majer, DC Nickle, JM Rommens, KM Popov, RA Harris, NL Riebow, J Xia, LC Tsui, C Bogardus, M Prochazka. Cloning and characterization of PDK4 on 7q21.3 encoding a fourth pyruvate dehydrogenase kinase isoenzyme in human. J Biol Chem 271:22376–22382, 1996. 52. T Katsube, S Nomoto, S Togashi, R Ueda, M Kobayashi, M Takahisa. cDNA sequence and expression of a gene encoding a pyruvate dehydrogenase kinase homolog of Drosophila melanogaster. DNA Cell Biol 16:335–339, 1997. 53. W Chen, X Huang, PR Komuniecki, R Komuniecki. Molecular cloning, functional expression, and characterization of pyruvate dehydrogenase kinase from anaerobic muscle of the parasitic nematode Ascaris suum. Arch Biochem Biophys 353:181–189, 1998. 54. JJ Thelen, MG Muszynski, JA Miernyk, DD Randall. Molecular analysis of two pyruvate dehydrogenase kinases from maize. J Biol Chem 273:26618– 26623, 1998. 55. JJ Thelen, JA Miernyk, DD Randall. Pyruvate dehydrogenase kinase from Arabidopsis thaliana: a protein histidine kinase that phosphorylates serine residues. Biochem J 349:195–201, 2000. 56. MM Bowker-Kinley, WI Davis, P Wu, RA Harris, KM Popov. Evidence for existence of tissue-speciﬁc regulation of the mammalian pyruvate dehydrogenase complex. Biochem J 329:191–196, 1998. 57. MC Sugden, HS Lall, RA Harris, MJ Holness. Selective modiﬁcation of the pyruvate dehydrogenase kinase isoform proﬁle in skeletal muscle in hyperthyroidism: implications for the regulatory impact of glucose on fatty acid oxidation. J Endocrinol 167:339–345, 2000. 58. B Huang, P Wu, MM Bowker-Kinley, RA Harris. Regulation of pyruvate dehydrogenase kinase expression by peroxisome proliferator-activated receptor-alpha ligands, glucocorticoids, and insulin. Diabetes 51:276–283, 2002.

508

Patel and Korotchkina

59. MJ Holness, K Bulmer, GF Gibbons, MC Sugden. Up-regulation of pyruvate dehydrogenase kinase isoform 4 (PDK4) protein expression in oxidative skeletal muscle does not require the obligatory participation of peroxisome-proliferator-activated receptor alpha (PPARalpha). Biochem J 366:839–846, 2002. 60. D Yang, X Gong, A Yakhnin, TE Roche. Requirements for the adaptor protein role of dihydrolipoyl acetyltransferase in the up-regulated function of the pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase. J Biol Chem 273:14130–14137, 1998.

28 The Complexity of Single-Gene Disorders: Lessons from Maple Syrup Urine Disease and Thiamine Responsiveness Dean J. Danner, Eric A. Muller, and Andrea Kasinski Emory University School of Medicine, Atlanta, Georgia, U.S.A.

I. HISTORY OF MAPLE SYRUP URINE DISEASE It has been almost 50 years since the ﬁrst description appeared for a disease that was lethal in infancy, that ran in families, and for which aﬀected individuals had a sweet odor associated with their urine (1). Dr. Menkes likened the odor to maple syrup and hence coined the name maple syrup urine disease (MSUD). The expression pattern for this condition ﬁts the classic deﬁnition of an ‘‘inborn error of metabolism,’’ as deﬁned by Sir Archibald Garrod, since it adhered to Mendelian genetic predictions for an autosomal recessive trait. In subsequent studies Menkes identiﬁed the compounds that produced the odor as the branched-chain amino acids (BCAA) and their transaminated a-ketoacids (BCKA) (2). In a series of reports following these initial studies, Dancis and colleagues showed that the defect resulted from the inability to decarboxylate the BCKA (3–6). This enzyme activity resided in the mitochondrion and was shown to be present in all mammalian cells with this organelle. The similarity of substrates and products to the characterized pyruvate dehydrogenase complex (PDC) allowed the study of the BCKA enzyme to follow a similar path of study (7,8). The enzyme was named 509

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branched-chain a-ketoacid dehydrogenase (BCKD). When it was isolated and puriﬁed to homogeneity, BCKD was found to convert all three BCKA to their acyl-CoA esters (9,10). The properties of the BCKD complex and its position in the BCAA metabolic pathway make it the committed step in BCAA catabolism. Although it was established that impaired function of BCKD caused MSUD, the ﬁrst clearly deﬁned molecular basis for the dysfunction was not demonstrated until 1985. Western blots of mitochondrial proteins from patient-derived cell lines showed an antigenic absence of one of the subunits of the complex (11). Shortly thereafter the ﬁrst cDNA clone for this missing subunit was isolated. This initiated an ongoing quest to deﬁne the molecular genetic basis and nucleotide changes in the mutant alleles that cause MSUD (12,13). II. BCKD COMPLEX Studies with the puriﬁed BCKD complex demonstrated that catalytic activity resided in four protein subunits (9,10). Enzyme activity depended on the presence of four cofactors (thiamine pyrophosphate [TPP], divalent magnesium, coenzyme A [CoASH], and NAD+), with the formation of three products (CO2, branched-chain acyl-CoA, and NADH) in a 1:1:1 stoichiometry (14). Two proteins, E1a and E1h, form a tetrameric structure and use TPP/Mg2+ to catalyze the decarboxylation of the BCKA (15). The residual branched-chain acyl group is transferred to CoASH by the action of the acyltransferase (E2) component. E2 contains a covalently bound lipoic acid that forms a thiol ester intermediate with the branched-chan acyl moiety in the path to forming the branched-chain CoA ester. Reoxidation of the reduced lipoate occurs by the action of the ﬂavoprotein lipoamide dehydrogenase [E3] and the ultimate reduction of NAD+. This E3 protein also functions with the pyruvate and a-ketoglutarate dehydrogenase complexes and is a component of the glycine cleavage complex (16,17). The physical arrangement of the subunits in the matrix of mitochondria has been speculated to involve a core of 24 E2 proteins surrounded by 12 E1 tetramers and 6 E3 dimers (16). It has not been clearly deﬁned whether the dynamics of this physical arrangements is involved in the regulation of catalytic activity, although reports in the literature suggest that the relative concentrations of the individual subunits are subject to change in response to diet and hormones in some tissues (18–20). III. NEED FOR BCKD REGULATION Since the BCKD complex is present in all cells with mitochondria, the activity state must be controlled to preserve or degrade the BCAA as needed for

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proper cellular function. BCKD activity commits the BCAA to their catabolic fate and loss from a cell. However, this catabolism contributes little to the energy production of cells since the absolute amount of NADH produced is small relative to that produced by carbohydrate metabolism. Cells are able to sense when the concentration of leucine drops below a critical threshold, and this accelerates protein degradation, especially in muscle (21). Uncontrolled protein loss from muscle is known as cachexia, a consequence that occurs in late stages of diseases such as diabetes, cancer, and AIDS and can be a component of the normal process of aging. The exact involvement of BCKD in this process is not known (22–24). In brain tissue, the BCAA serve as the major source of nitrogen for the formation of glutamate (25). This preference is due at least in part to the fact that BCAA cross the blood–brain barrier more readily than other amino acids. During reverse transamination the BCKA will accept nitrogen from glutamate, serving to decrease the glutamate concentration and its signaling eﬀects. Therefore the activity state of BCKD in glial and neuronal cells must be carefully controlled to maintain the necessary concentrations of the BCAA and BCKA in each cell type. Imbalance in these reactions can lead to seizures and epileptic conditions.

Figure 1 Branched-chain amino acid uptake, metabolism, and functional consequences known to be involved in different cell types.

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Another role for leucine is to stimulate insulin release from the h-cell of the pancreas, and it trails only glucose in this function (26–28). Thus, it again becomes important for the concentration of leucine to be tightly controlled in these cells. If leucine were to remain in high concentration, there is the risk for unneeded release of insulin. The activity state of BCKD must be adequate to ensure that leucine concentration in the h-cells remains at a concentration commensurate with the need for insulin release. The contribution of BCKD dysfunction or lack of controlled activity state in the mechanism of diabetes is not known. Individuals with MSUD may be at an increased risk for developing diabetes as they age, a condition that merits close monitoring. The various roles of BCAA and their metabolism are illustrated as a cartoon in Figure 1.

IV. REGULATION OF BCKD ACTIVITY STATE BY THE COMPLEX-SPECIFIC KINASE Regulation of the activity state of the complex is known to occur through phosphorylation of the E1a subunit on two serine residues, S337 and S347 (29). Addition of these phosphates occurs from the action of a complex-speciﬁc kinase (BCKD-kinase) (30–32). Characterization of a putative BCKD-phosphatase remains to be done, since the isolation of this protein has not been repeated and a gene encoding a protein with this function has not been found (33,34). From crystalographic studies of the E1 tetramer it was shown that phosphate attachment at S337 blocks the entry of the ketoacid to the substratebinding site and may also interfere with TPP binding (15). No importance for phosphorylation of S347 has been found (35). In the fed state, skeletal muscle has a high expression level of the BCKDkinase, with a concomitant low activity state of the BCKD complex. In contrast, liver mitochondria contain little of the BCKD-kinase protein, and thus most of the BCKD complex is catalytically active. Mitochondria from tissues such as brain, kidney, and heart contain levels of the BCKD-kinase protein that is intermediate between these two extremes (36,37). Dietary protein and certain hormones have been shown to alter the BCKD-kinase protein that is present in liver and skeletal muscle, but the eﬀect of these conditions on kinase expression in other tissues has not been reported (38–40). The majority of these studies were done with the laboratory rat as a model. Studies have also been done in cultured cells, demonstrating that the absence of BCAA in the culture medium increases kinase expression with a coupled decrease in BCKD activity state (41,42). Doering et al. demonstrated that the absence of medium BCAA induced the speciﬁc translation of existing message for BCKD-kinase without a change in gene transcription (42). None

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of these studies examined the eﬀect of thiamine or TPP on BCKD activity or kinase expression.

V. CLINICAL PHENOTYPE AND MUTATIONS CAUSING MSUD A. Phenotypes The genotype and BCKD activity are not clearly deﬁned for the majority of individuals with MSUD. Clinical diagnosis is most often made on plasma BCAA values alone. For an autosomal recessive trait, heterozygote parents should express 50% of normal enzyme activity, but in reality many obligate heterozygote parents are found to have BCKD activity near that found in wild-type cells. For this reason it is not possible to ascertain carrier status on BCKD activity measurements. This discrepancy is likely due to the multiprotein nature of the complex and the involvement of complex assembly in determining overall activity. We know little of how expression of other genes may inﬂuence BCKD activity, but it is well established that single-gene traits exhibit a complex and varied phenotype (43,44). Even siblings with MSUD can show diﬀerences in protein tolerance and susceptibility to metabolic decompensation, attesting to the inﬂuence of one’s genetic background on the phenotype. Prior to the late 1960s, when the use of protein-modiﬁed diets was begun as treatment, individuals with MSUD died in infancy, and death was often attributed to sudden infant death syndrome. As the knowledge of the disease grew, a clinical classiﬁcation was used based on the time after birth when clinical signs ﬁrst appeared. These signs include lethargy, seizures, coma, and possible death accompanied by elevated plasma concentration of BCAA and BCKA. Individuals presenting in the ﬁrst two weeks postnatal were termed ‘‘classic’’ and considered the most severely aﬀected. BCKD activity is usually below 2–5% of normal when assayed in cells cultured from these individuals. It appears that plasma leucine concentrations are most damaging and have been reported as high as 5000 lM relative to the normal value of near 100 lM (45). A second classiﬁcation was given to patients that presented in the age range of 1–8 years and followed a less severe clinical course. These individuals were labeled as ‘‘intermediate’’ or ‘‘intermittent’’ MSUD, with BCKD activity in the range of 5–30% of normal. For the majority of individuals in this group the enzyme assays were done without regard to the phosphorylation state of the wild-type enzyme complex. Therefore, the percent normal may have been compared with a partially active wild-type complex, making the percent of BCKD that is active in aﬀected individuals appear to be higher

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than the actual values. A systematic reassessment of the BCKD activity in the ‘‘intermediate/intermittent’’ group has not been done. BCKD activity must be reported relative to a fully active native complex in a culture cell derived from the same tissue that was used for patient analysis. Another classiﬁcation used clinically for MSUD, ‘‘thiamine-responsive,’’ is of direct interest to the topic of this chapter. The term was ﬁrst used to describe a single patient who showed a clinical improvement when her protein-modiﬁed diet was supplemented with pharmacologic levels of thiamine (46). A second report, 15 years later, by the same authors questioned their earlier report and suggested their patient did not demonstrate a continued long-term response to thiamine (47). Other reports support the existence of these thiamine-responsive patients, but no long-term surveillance studies have appeared (48–50). Attempts have been made to deﬁne a mechanism for the improved BCKD activity when the protein-modiﬁed diet is supplemented with pharmacologic thiamine, but none have been clearly elucidated (51,52). It is possible that multiple mechanisms are working with different mutant genotypes and genetic backgrounds. Too few patients with MSUD exist in any one genotype to accurately deﬁne the mechanism of thiamine response with available methods. The largest population of individuals with the same genotype is the Mennonites, and the standard dietary treatment used for the majority of these individuals does not include the thiamine supplement (53). A controlled study to determine the value of thiamine supplementation in this group has not been reported. Since there are no reports of thiamine toxicity, supplementing the protein-modiﬁed diet of all patients with MSUD is common practice with some treatment centers. It should be pointed out, however, that tailoring the protein-modiﬁed diet to each individual is the prudent approach used by most treatment centers. B. Mutations Some variability in phenotype can be expected, since BCKD is a multiprotein complex and dysfunction can result from inherited mutations in any of the genes that encode these proteins. The three genes unique to the BCKD complex, which harbor causative mutations for MSUD, are depicted in Figure 2. The amino acid sequences of these proteins are highly conserved throughout evolution. Antisera raised against the bovine proteins recognize these antigens in all mammalian cells as well as proteins with the same electrophoretic mobility in the invertebrate C. elegans. Survey of the C. elegans genome database reveals BCKD subunit orthologs, but BCKD activity has not been demonstrated. Based on the degree of conservation of the amino acid sequences it folows that nonconservative amino acids substitutions throughout these proteins result in dysfunction. There have been over 50 missense mutations

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Figure 2 Gene structure and chromosome location of the catalytic subunits unique to the BCKD complex and known to harbor mutations that cause MSUD.

described among the three genes to date (Fig. 3). In most families the aﬀected individual will have received a diﬀerent mutant allele from each parent in the same gene to result in MSUD. Consanguinity has resulted in some individuals being homozygous for a single mutation (54). MSUD in the Mennonites is due to a common allele that changes tyrosine 483 to asparagine (Y438N) in the E1a protein. An amino acid substitution can alter substrate- and/or cofactor-binding sites or change the ability of the protein to associate with the other proteins in the complex. Ævarson et al. have solved the crystal structure for the E1 a2h2 tetramer and have made predictions for consequences of known amino acid substitutions found in MSUD. Their analyses predicted disruption of TPP binding for the R159W and R265W substitutions that occur in E1a (15). The study further predicted that the T211M amino acid mutation causes dysfunction due to interference with potassium binding. It is interesting to note that during the puriﬁcation and characterization of the BCKD complex, both reports made reference to the importance of using potassium phosphate buﬀers to maintain activity during the isolation. When sodium phosphate buﬀers were used, activity was lost (9,10). The described role for K+ in the tetramer oﬀers an explanation for earlier observations regarding the need for potassium buﬀers. A number of other amino acid substitutions were reported to alter protein–protein interactions; the most noted is the one common allele

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Figure 3 Missense mutations and the exon position that result in dysfunction of the BCKD complex.

found in the Mennonite population. This Y438N substitution in the carboxy terminal end of the E1a subunit prevents its association with the E1h subunit, and therefore the tetrameric structure is not formed (15,55). C. Thiamine-Responsive MSUD Mutations other than missense or nonsense within the exons are known, and a total of more than 80 disease-causing alleles have been deﬁned. Some nucleic acid changes result in no detectable protein for a subunit. This has been shown to result from a genomic deletion, in at least one case. The absence of protein can also be the result of decreased message stability, improper processing, or failure of translation (11,56,57). It is also possible that protein stability could be decreased and result in the observed absence. Individuals with the most clearly deﬁned response to thiamine supplementation have the genotype that results in the null phenotype for E2 protein expression. Apparently the E1 tetramer forms normally, even in the absence of E2. Western blots of mitochondrial proteins in cells prepared from individuals with this genotype show the antigenic presence of both E1a and E1h in amounts equal to that observed in cells from individuals with normal BCKD activity (52,58). One proposed

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Figure 4 Proposed mechanism for thiamine-responsive MSUD. The E1 a2h2 tetramer is illustrated with and without the TPP cofactor. BCacyl represents the branchedchain acyl group after decarboxylation has occurred.

mechanism holds that the TPP–branched-chain acyl moiety is displaced from E1 and this conjugate is excreted. Since the dietary supplement of thiamine allows for excess TPP to be available, the E1 tetramer binds another molecule of cofactor and can undergo another round of decarboxylation of the BCKA (Fig. 4). Another mechanism proposed is based on the mutations aﬀecting the TPP-binding site (51). It is likely that more than one mechanism may explain thiamine-responsive MSUD, since a wide variety of mutations alter the function of BCKD. Skin and/or blood cells cultured from these individuals have not shown a BCKD response to TPP, so the response mechanism was not elucidated. One explanation, among several, may be that only selected tissues will respond (52). Finding a speciﬁc tissue in which the thiamine supplementation augments BCKD activity will target those tissues for directed gene therapy and improved treatment protocols in the future. VI. ROLE OF TPP IN NORMAL BCKD FUNCTION TPP binding to the functional BCKD complex induces a conformational change in the complex, as demonstrated by CD/ORD spectroscopy (59). This conformational change in subunit association results in increased stability of the complex against proteolytic degradation or could inﬂuence BCKD-kinase binding to the complex (60). It has been reported that TPP can block the

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ability of BCKD-kinase to inactivate BCKD, but the mechanism for this inhibition has not been described (61). Additional studies for the eﬀect on TPP on the kinase have not been reported. VII. MODEL SYSTEMS Neuronal dysfunction as a consequence of elevated plasma BCAA concentrations is poorly understood. The naturally occurring bovine MSUD model has provided some understanding; however, too few of these animals exist for in-depth analyses (62). Since brain tissue from humans with MSUD is not readily available for study, we developed a model system using PC12 rat phenochromocytoma cells that express the rtTA transactivator protein (63). A plasmid with the mouse cDNA for the BCKD-kinase inserted behind the tetracycline response elements (TRE), constructed by Doering et al., was transfected into these rtTA-expressing PC12 cells (64). Cells harboring the kinase transgene were clonally selected and tested for the ability of doxycycline (DOX) to stimulate expression of the kinase transgene (Fig. 5). Addition of DOX to the medium was able to induce expression of the transgene BCKDkinase protein in a time-dependent manner. The high expression of this transgene kinase resulted in a near 80% reduction in total BCKD activity (11.5 F 2.7 vs 2.7 F 0.1 nmol CO2/mg protein/3 h). The amount of BCKD that is active in these cells approximates that found in cells from individuals with MSUD. Therefore these DOX-treated cells will serve as a model for studying the consequences of loss of BCKD in neuronal tissue. These cells were used to determine whether thiamine supplementation of the culture medium would result in a blocking of BCKD-kinase phosphorylation of the E1a subunit. An unexpected ﬁnding was that medium supplemented with 100 mg thiamine per liter resulted in stimulation of total BCKD activity in the cells. Standard DMEM used to culture these cells does not contain thiamine and therefore could explain the eﬀect of thiamine on enzyme activity. The activity state of BCKD in cells treated with thiamin was found to be 50%. The activity state after DOX addition was lowered but

Figure 5 Western blot of mitochondrial proteins from PC12 cells that have been treated with 1 Ag/mL of DOX for the indicated time using antiserum against BCKDkinase. Each lane contains 20 Ag of total mitochondrial protein.

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Table 1 Thiamine Eﬀect on BCKD-Kinase Changing the Activity State of BCKD (Percent of BCKD That Is Active)a Thiamine 100 mg/L

No DOX Plus DOX

+

100 38

51 37

a

Thiamine was added to the medium 24 h prior to the assay, and DOX was added 4 h prior to assay. The data presented represent one of two independent observations.

unchanged after thiamine addition (Table 1.) These results demonstrate the utility of the constructed PC12 cells to investigate neuronal cell function under a variety of activity states for BCKD. Of special interest will be the role of thiamine in BCKD function and BCKD-kinase expression.

REFERENCES 1.

2. 3. 4. 5.

6. 7.

8.

JH Menkes, PL Hurst, JM Craig. A new syndrome: progressive familial infantile cerebral dysfunction associated with an unusual urinary substance. Pediatrics 14:462–467, 1954. JH Menkes. Maple syrup disease Isolation and identiﬁcation of organic acids in the urine. Pediatrics 23:348–353, 1959. J Dancis, J Hutzler, M Levitz. Metabolism of the white blood cells in maple syrup urine disease. Biochim Biophys Acta 43:342–343, 1960. J Dancis, J Hutzler, M Levitz. Tissue distribution of branched-chain ketoacid decarboxylase. Biochim Biophys Acta 52:60–64, 1961. J Dancis, J Hutzler, RP Cox. Enzyme defect in skin ﬁbroblasts in intermittent branched-chain ketonuria and in maple syrup urine disease. Biochem Med 2: 407–411, 1969. J Dancis, J Hutzler, SE Snyderman, RP Cox. Enzyme activity in classical and variant forms of maple syrup urine disease. J Pediatr 81:312–320, 1972. T Hayakawa, M Koike. Mammalian alpha-keto acid dehydrogenase complexes. 3 Resolution and reconstitution of the pig heart pyruvate dehydrogenase complex. J Biol Chem 242:1356–1358, 1967. F Hucho, DD Randall, TE Roche, MW Burgett, JW Pelley, LJ Reed. a-Keto acid dehydrogenase complexes. XVII. Kinetic and regulatory properties of pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase from bovine kidney and heart. Arch Biochem Biophys 151:328–340, 1972.

520 9.

10.

11.

12.

13.

14.

15.

16. 17. 18. 19.

20.

21. 22.

23. 24.

Danner et al. DJ Danner, SK Lemmon, JC Besharse, LJ Elsas. Puriﬁcation and characterization of branched-chain a-ketoacid dehydrogenase from bovine liver mitochondria. J Biol Chem 254:5522–5526, 1979. FH Pettit, SJ Yeaman, LJ Reed. Puriﬁcation and characterization of branchedchain a-ketoacid dehydrogenase complex of bovine kidney. Proc Natl Acad Sci USA 75:4881–4886, 1978. DJ Danner, N Armstrong, SC Heﬀelﬁnger, ET Sewell, JH Priest, LJ Elsas. Absence of branched-chain acyl-transferase as a cause of maple syrup urine disease. J Clin Invest 75:858–860, 1985. S Litwer, DJ Danner. Identiﬁcation of a cDNA clone in Egt11 for the transacylase component of branched-chain ketoacid dehydrogenase. Biochem Biophys Res Comm 131:961–967, 1985. DT Chuang, VE Shih. Maple syrup urine disease (branched-chain ketoaciduria). In: CR Scriver, AL Beaudet, WS Sly, D Valle, eds. The Metabolic and Molecular Basis of Inherited Disease. New York: McGraw-Hill, 2:1971–2005, 2001. SC Heﬀelﬁnger, ET Sewell, DJ Danner. Identiﬁcation of speciﬁc subunits of highly puriﬁed bovine liver branched-chain ketoacid dehydrogenase. Biochemistry 22:5519–5522, 1983. A Ævarsson, JL Chuang, RM Wynn, S Turley, DT Chuang, WGJ Hol. Crystal structure of human branched-chain a-ketoacid dehydrogenase and the molecular basis of multienzyme complex deﬁciency in maple syrup urine disease. Structure Fold Des 8:277–291, 2000. LJ Reed, ML Hackert. Structure-function relationships in dihydrolipoamide acyltransferases. J Biol Chem 265:8971–8974, 1990. SJ Yeaman. The 2-oxo acid dehydrogenase complexes: recent advances. Biochem J 257:625–632, 1989. PA Costeas, JM Chinsky. Glucocorticoid regulation of branched-chain a-ketoacid dehydrogenase E2 subunit gene expression. Biochem J 347:449–457, 2000. PA Costeas, JM Chinsky. Eﬀects of insulin on the regulation of branched-chain a-ketoacid dehydrogenase E1a subunit gene expression. Biochem J 318:85–92, 1996. Y Zhao, KM Popov, Y Shimomura, NY Kedishvili, J Jaskiewicz, MJ Kuntz, J Zhang, B Zhang, RA Harris. Eﬀect of dietary protein on the liver content and subunit composition of the branched-chain a-ketoacid dehydrogenase complex. Arch Biochem Biophys 308:446–453, 1994. SR Kimball, LS Jeﬀerson. Regulation of protein synthesis by branched chain amino acids. Curr Opin Clin Nutr Metab Care 4:39–43, 2001. DC Hunter, M Weintraub, GL Blackburn, BR Bistrian. Branched-chain amino acids as the protein component of parenteral nutrition in cancer cachexia. Br J Surg 76:149–153, 1989. PWT Pisters, MF Brennan. Amino acid metabolism in human cancer cachexia. Ann Rev Nutr 10:107–132, 1990. PWT Pisters, DB Pearlstone. Protein and amino acid metabolism in cancer cachexia: investigative techniques and therapeutic interventions. Crit Rev Clin Lab Sci 30:223–272, 1993.

MSUD and Thiamine Responsiveness

521

25. M Yudkoﬀ. Brain metabolism of branched-chain amino acids. Glia 21:92–98, 1997. 26. WJ Malaisse, JC Hutton, AR Carpinelli, A Herchuelz, A Sener. The stimulussecretion coupling of amino acid-induced insulin release: metabolism and cationic eﬀects of leucine. Diabetes 29:431–437, 1980. 27. MJ MacDonald, DI McKenzie, JH Kaysen, TM Walker, SM Moran, LA Towle, HC Towle. Glucose regulates leucine-induced insulin release and the expression of the branched-chain ketoacid dehydrogenase E1a subunit gene in pancreatic islets. J Biol Chem 266:1335–1340, 1991. 28. JC Floyd, SS Fajans, RF Knopf, JW Conn. Evidence that insulin release is the mechanism for experimentally-induced leucine hypoglycemia in man. J Clin Invest 42:1714–1719, 1963. 29. KG Cook, R Lawson, SJ Yeaman. Multi-site phosphorylation of bovine kidney branched-chain 2-oxoacid dehydrogenase complex. FEBS Lett 157:59–62, 1983. 30. R Odessey. Puriﬁcation of rat kidney branched-chain oxo acid dehydrogenase complex with endogenous kinase activity. Biochem J 204:353–356, 1982. 31. R Odessey. Direct evidence for the inactivation of branched-chain oxo-acid dehydrogenase by enzyme phosphorylation. FEBS Lett 121:306–308, 1980. 32. KM Popov, Y Shimomura, RA Harris. Puriﬁcation and comparative study of the kinases speciﬁc for branched-chain a-ketoacid dehydrogenase and pyruvate dehydrogenase. Prot Exp Purif 2:278–286, 1991. 33. Z Damuni, ML Merryﬁeld, JS Humphreys, LJ Reed. Puriﬁcation and properties of branched-chain a-ketoacid dehyrogenase phosphatase from bovine kidney. Proc Natl Acad Sci USA 81:4335–4338, 1984. 34. Z Damuni, LJ Reed. Puriﬁcation and properties of the catalytic subunit of the branched-chain a-ketoacid dehydrogenase phosphatase from bovine kidney mitochondria. J Biol Chem 262:5129–5132, 1987. 35. JW Hawes, RJ Schnepf, AE Jenkins, Y Shimomura, KM Popov, RA Harris. Roles of amino acid residues surronding phosphorylation site 1 of branchedchain a-ketoacid dehydrogenase (BCKDH) in catalysis and phorphorylation site recognition by BCKDH kinase. J Biol Chem 270:31071–31076, 1995. 36. GW Goodwin, W Gibboney, R Paxton, RA Harris, JA Lemons. Activities of branched-chain amino acid aminotransferase and branched-chain 2-oxo acid dehydrogenase complex in tissues of maternal and fetal sheep. Biochem J 242: 305–308, 1987. 37. AJM Wagenmakers, JTG Schepens, JAM Veldhuizen, JH Veerkamp. The activity state of the branched-chain 2-oxoacid dehydrogenase complex in rat tissues. Biochem J 220:273–281, 1984. 38. KP Block, RP Aftring, MG Buse. Regulation of rat liver branched-chain aketoacid dehydrogenase activity by meal frequency and dietary protein. J Nutr 120:793–799, 1990. 39. KP Block, WB Richmond, WB Mehard, MG Buse. Glucocorticoid-mediated activation of muscle branched-chain a-keto acid dehydrogenase in vivo. Am J Physiol 252:E396–E407, 1987. 40. Y Shimomura, M Obayashi, T Murakami, RA Harris. Regulation of branchedchain amino acid catabolism: nutritional and hormonal regulation of activity

522

41.

42.

43.

44. 45. 46. 47.

48. 49.

50. 51.

52.

53.

54.

55.

56.

Danner et al. and expression of the branched chain a-keto acid dehydrogenase kinase. Curr Opin Clin Nutr Metab Care 4:419–423, 2001. M Beggs, JM Shaw, PJ Randle. Longer-term regulation of branched-chain-2oxoacid dehydrogenase complex studied in rat hepatocytes in culture. Biochem J 257:271–275, 1989. CB Doering, DJ Danner. Amino acid deprivation induces translation of branched-chain a-ketoacid dehydrogenase kinase. Am J Physiol 279:C1587– C1594, 2000. KM Dipple, ERB McCabe. Phenotypes of patients with ‘‘simple’’ mendelian disorders are complex traits: thresholds, modiﬁers, and systems dynamics. Am J Hum Genet 66:1729–1735, 2000. CR Scriver, PJ Waters. Monogenic traits are not simple. Trends Genet 15:267– 272, 1999. F Peinemann, DJ Danner. Maple syrup urine disease 1954 to 1993. J Inherit Metab Dis 17:3–15, 1994. CR Scriver, CL Clow, S Mackenzie, E Delvin. Thiamine-responsive maplesyrup-urine disease. Lancet i:310–312, 1971. CR Scriver, CL Clow, H George. So-called thiamin-responsive maple syrup urine disease: 15-year follow-up of the original patient. J Pediatr 107:763–765, 1985. M Duran, SK Wadman. Thiamine-responsive inborn errors of metabolism. J Inherit Metab Dis 8:70–75, 1985. PM Fernhoﬀ, D Lubitz, DJ Danner, PP Dembure, HP Schwarz, R Hillman, DM Bier, LJ Elsas. Thiamine-responsive maple syrup urine disease. Pediat Res 19:1011–1016, 1985. LJ Elsas, DJ Danner. The role of thiamine in maple syrup urine disease. Ann NY Acad Sci 378:404, 1982. DT Chuang, LS Ku, RP Cox. Thiamin-responsive maple-syrup-urine disease: decreased aﬃnity of the mutant branched-chain a-keto acid dehydrogenase for a-ketoisovalerate and thiamin pyrophosphate. Proc Natl Acad Sci USA 79: 3300–3304, 1982. NP Ellerine, WJ Herring, LJ Elsas, MC McKean, PD Klein, DJ Danner. Thiamin-responsive maple syrup urine disease in a patient antigenically missing dihydrolipoamide acyltransferase. Biochem Med Metab Biol 49:363–374, 1993. DH Morton, KA Strauss, DL Robinson, EG Puﬀenberger, RI Kelley. Diagnosis and treatment of maple syrup disease: a study of 36 patients. Pediatrics 109:999–1008, 2002. A Dursun, M Henneke, K Ozgul, J Gartner, T Coskun, A Tokatli, HS Demirkol, M Demirkol, U Wendel, I Ozalp. Maple syrup urine disease: mutation analysis in Turkish patients. J Inherit Metab Dis 25:89–97, 2002. CR Fisher, JL Chuang, RP Cox, CW Fisher, RA Star, DT Chuang. Maple syrup urine disease in Mennonites: evidence that the Y393N mutation in E1alpha impedes assembly of the E1 component of branched-chain a-ketoacid dehydrogenase complex. J Clin Invest 88:1034–1037, 1991. WJ Herring, S Litwer, JL Weber, DJ Danner. Molecular genetic basis of maple

MSUD and Thiamine Responsiveness

57.

58.

59.

60.

61. 62.

63.

64.

523

syrup urine disease in a family with two defective alleles for branched-chain acyltransferase and localization of the gene to human chromosome 1. Am J Hum Genet 48:342–350, 1991. WJ Herring, M McKean, N Dracopoli, DJ Danner. Branched-chain acyltransferase absence due to an ALU-based genomic deletion allele and an exon skipping allele in a compound heterozygote proband expressing maple syrup urine disease. Biochim Biophys Acta 1138:236–242, 1992. S Litwer, WJ Herring, DJ Danner. Reversion of maple syrup urine disease phenotype of impaired branched-chain a-ketoacid dehydrogenase complex activity in ﬁbroblasts from an aﬀected child. J Biol Chem 264:14597–14600, 1989. DJ Danner, SK Lemmon, LJ Elsas. Stabilization of mammalian liver branchedchain a-ketoacid dehydrogenase by thiamin pyrophosphate. Arch Biochem Biophys 202:23–28, 1980. SC Heﬀelﬁnger, ET Sewell, LJ Elsas, DJ Danner. Direct physical evidence for stabilization of branched-chain ketoacid dehydrogenase by thiamin pyrophosphate. Am J Hum Genet 36:802–807, 1984. KS Lau, HR Fatania, PJ Randle. Regulation of the branched-chain 2-oxoacid dehydrogenase kinase reaction. FEBS Lett 144:57–62, 1982. HA Harper, JA Dennis, PJ Healy, GK Brown. Maple syrup urine disease in calves: a clinical, pathological and biochemical study. Aust Vet J 66:46–49, 1989. N Jha, O Jurma, G Lalli, Y Liu, EH Pettus, JT Greenamyre, RM Liu, HJ A, JK A. Glutathione depletion in PC12 results in selective inhibition of mitochondrial complex I activity. Implications for Parkinson’s disease. J Biol Chem 275:26096–26101, 2000. CB Doering, IR Williams, DJ Danner. Small-molecule regulation of BCKD kinase expression: a tool for controlling cellular BCAA metabolism. Metabol Eng 2:349–356, 2000.

29 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 Faculty of Medicine, Katholieke Universiteit Leuven, Leuven, Belgium

I. INTRODUCTION During our research on a-oxidation of 3-methyl-branched fatty acids, we recently discovered a new enzyme, 2-hydroxyphytanoyl-CoA lyase, catalyzing the third step of the proposed pathway and found it to be thiamine dependent (1). Until then, no thiamine dependence of any part of the alphaoxidation pathway had been suspected. Because alpha-oxidation is a new ﬁeld in thiamine biochemistry, this chapter starts with an overview of a-oxidation of 3-methyl-branched fatty acids and the pathology caused by its deﬁciency. Then the thiamine-dependent enzyme 2-hydroxyphytanoyl-CoA lyase is described in more detail. The 3-methyl-branched fatty acid best known and most important to the human species is phytanic acid (3,7,11,15-tetramethylhexadecanoic acid), which is derived from phytol, the isoprenoid side chain of chlorophyll. Several human pathologies are linked with the accumulation of phytanic acid, the best known of which is adult Refsum disease (ARD) (2,3). 525

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Figure 1 Chemical structures of chlorophyll, phytol, and phytanic acid (3,7,11,15tetramethylhexadecanoic acid).

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Because chlorophyll-bound phytol cannot be metabolized by humans and free phytol is present in only minimal quantities in food, the phytanic acid present in the human body is provided mostly from external sources (Fig. 1). Ruminants ingest large amounts of chlorophyll, from which phytol is efﬁciently cleaved oﬀ by bacteria in the gastrointestinal tract. Phytol is subsequently converted to phytanic acid, which is deposited in fat tissues and in milk, the major sources of phytanic acid for humans. The classic catabolic pathway by which fatty acids are degraded is h-oxidation. Both a mitochondrial and a peroxisomal h-oxidation sequence are known, each with a more or less deﬁned substrate speciﬁcity. Phytanic acid and other 3-methylbranched fatty acids cannot undergo a-oxidation because the 3-methyl group prevents the formation of a 3-keto substituent in the dehydrogenation step. For this reason 3-methyl-branched fatty acids ﬁrst undergo a-oxidation, a peroxisomal process in which the fatty acid is shortened by only one carbon atom. In the case of phytanic acid, a-oxidation results in the 2-methylbranched pristanic acid (2,8,10,14-tetramethylpentadecanoic acid), which is then degraded to 4,8-dimethyl-nonanoic acid via peroxisomal h-oxidation. The dimethyl fatty acid is then degraded further via mitochondrial hoxidation. Peroxisomes are subcellular organelles involved in a number of anabolic (e.g., plasmalogen synthesis) and catabolic processes, and they are indispensable for the a-oxidation of long 3-methyl-branched fatty acids and for the h-oxidation of very long-chain fatty acids, long 2-methyl-branched fatty acids, and the side chains of bile acid intermediates (4). Peroxisomal enzymes are synthesized in the cytosol and are posttranslationally directed to the peroxisome. To make them ﬁnd their way to the organelle, they contain a series of conserved amino acids, or the so-called peroxisome-targeting signals (PTSs). Two classes of these topogenic sequences have been described: PTS1, a carboxy-terminal tripeptide, and PTS2, an amino-terminal nonapeptide (5). II. A-OXIDATION OF 3-METHYL-BRANCHED FATTY ACIDS Only in the last decade have most aspects of a-oxidation been unraveled (4). A major breakthrough was Poulos’ ﬁnding that in ﬁbroblasts a-oxidation of 3methyl-branched fatty acids generated not only CO2 but also formate (6). Not all questions are fully resolved, but the following scheme (Fig. 2) represents the current knowledge about this process. In a ﬁrst step, the 3-methylbranched fatty acid is activated to the corresponding CoA-ester by an acylCoA synthetase present in the peroxisomal membrane. It is not clear which synthetase is responsible for the activation step: a nonspeciﬁc long-chain fatty

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Figure 2 a-Oxidation of 3-methylbranched fatty acids. This scheme represents the a-oxidation pathway of 3-methyl-hexadecanoic acid [a valid substitute for studying phytanic acid metabolism (23)] and the stereochemical conﬁguration of the intermediates involved. The numbers indicate the enzymes catalyzing the diﬀerent steps: (1) acyl-CoA synthetase; (2) phytanoyl-CoA hydroxylase (PAHX); (3) 2-hydroxyphytanoyl-CoA lyase (HPCL); (4) formyl-CoA hydrolase; (5) aldehyde dehydrogenase; (6) acyl-CoA synthetase; and (7) 2-methylacyl-CoA racemase, responsible for the conversion of the R-2-methylacyl-CoA into the S-2-methylacyl-CoA, because only the S-isomer can undergo h-oxidation.

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acyl-CoA synthetase (7), a speciﬁc phytanoyl-CoA synthetase (8), or a very long-chain fatty acyl-CoA synthetase (9). In a second step, the 3-methyl-acyl-CoA is hydroxylated in position 2 by a dioxygenase, which is dependent on molecular O2, iron, 2-oxoglutarate, ascorbate, ATP/GTP, CoA, and Mg2+ (10–12). This 2-phytanoyl-CoA hydroxylase or PAHX contains a PTS2 signal (peroxisome-targeting signal 2) and is present in the peroxisomal matrix (13). The product of the reaction catalyzed by PAHX is a 2-hydroxy-3-methylacyl-CoA or, if phytanic acid is the substrate, 2-hydroxyphytanoyl-CoA. The 2-hydroxy-3-methylacyl-CoA is subsequently split in the peroxisomal matrix by 2-hydroxyphytanoyl-CoA lyase, which uses TPP as a cofactor (1). Products of this reaction are formyl-CoA (14) and a 2-methylbranched fatty aldehyde (15,16); the 2-methyl-branched fatty aldehyde is dehydrogenated by an NAD+-dependent aldehyde dehydrogenase, present in the peroxisomal matrix, to a 2-methyl-branched fatty acid (pristanic acid in the case of pristanal), which can be activated to the corresponding acyl-CoA ester. This CoA-ester can then enter the peroxisomal h-oxidation sequence. The formyl-CoA is hydrolyzed, mainly enzymatically, to formate in the peroxisome and converted to CO2 in the cytosol (14). III. DEFICIENT BREAKDOWN OF PHYTANIC ACID Patients with peroxisome biogenesis disorders or with a single enzymatic defect of a-oxidation accumulate phytanic acid in plasma and tissues. The most typical clinical picture of an isolated defect in phytanic acid breakdown is described as ‘‘adult Refsum disease,’’ or ARD (2,17). Only in the second or third decade does the gradual accumulation of phytanic acid in these patients produce distinct symptoms. Virtually all patients show retinitis pigmentosa (resulting in a typical picture when doing fundoscopy), night blindness, and anosmia (deﬁcient smelling sensation; 80% of ARD patients). In addition, polyneuropathy (60%), deafness (60%), ataxia (50%), and ichtyosis (20%) are quite common (for a review, see Ref. 17). In 30–40% of the patients, absence of a metacarpal or metatarsal is noted. A prerequisite for the diagnosis of ARD is the presence of an elevated serum level of phytanic acid (above 200 lM, whereas normal phytanic acid levels in serum are below 30 lM). The clinical spectrum of ARD can be attributed to diﬀerent molecular and genetic bases (18): (1) a defect at the level of PAHX, mapped to chromosome 10p13 (13,19); (2) a mutation in PEX7, speciﬁcally aﬀecting the peroxisomal import of PAHX (20); or (3) in atypical cases, a racemase deﬁciency (Ref. 21; see legend to Fig. 2). However, in some patients with the clinical syndrome of ARD none of these speciﬁc molecular defects could

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be found; the genetic basis of the disease in these patients awaits being deﬁned. IV. THE THIAMINE-DEPENDENT ENZYME OF THE A-OXIDATION PATHWAY, 2-HYDROXYPHYTANOYLCOA LYASE 2-Hydroxyphytanoyl-CoA lyase catalyzes the third step of the a-oxidation pathway, and was puriﬁed, characterized, sequenced, and cloned by Foulon et al. in 1999 (1). The discovery of this enzyme can be attributed to several important ﬁndings by diﬀerent groups. Up to 1993, only CO2, which was then considered the direct end product of a-oxidation, was measured when assaying a-oxidation, and—as a consequence—rates in broken cell fractions such as peroxisomes never equaled those in intact cells. In 1993 Poulos et al. (6) found that in ﬁbroblasts formate was also formed during a-oxidation of 3methyl-branched fatty acids. This was conﬁrmed for isolated hepatocytes in 1994 (22). It appeared that the formate/CO2 ratio increased as more disrupted cell systems were used (homogenates, subcellular fractions). In 1997, Croes et al. found that neither formate nor CO2 was the primary end product, but formyl-CoA (14). This ﬁnding led several authors to hypothesize a reaction mechanism in which the other product would be a 2-methyl aldehyde (or pristanal where phytanic acid was the substrate). Soon, the formation of a 2methyl aldehyde was demonstrated simultaneously by Croes et al. (15) and Verhoeven et al. (16). Foulon et al. used 2-OH-3-methylhexadecanoyl-CoA as substrate for studying the third reaction of the a-oxidation pathway; they measured formate (together with formyl-CoA, which is converted, partly enzymatically, to formate) as the primary reaction product. It had been shown before that 3methylhexadecanoic acid is metabolized in the same way as phytanic acid (23), and its use was validated as a substitute for the latter substrate when studying a-oxidation. The method for formate measurement, using mercuric acetate, was adapted from Yang, as described by Casteels et al. (22). Subcellular fractionation studies in rat liver demonstrated that the lyase colocalized with catalase in the peroxisomal fraction. Hence, isolation of the presumptive cleavage enzyme was started from the matrix protein fraction of isolated rat liver peroxisomes. The lyase was puriﬁed as a protein made up of four identical subunits of 63 kDa. Formyl-CoA and 2-methylpentadecanal (measured by GC analysis) were identiﬁed as reaction products when the enzyme puriﬁed in the presence of TPP (see later) was incubated with 2hydroxy-3-methylhexadecanoyl-CoA as the substrate. Quantitative measurements of both reaction products further conﬁrmed the stoichiometry of the cleavage step. Incubations in the presence of NAD+ [a cofactor for fatty aldehyde dehydrogenation (24)] did not alter the amount of formate and

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2-methylpentadecanal formed. Methylation of the hexane extracts of the incubation mixtures and subsequent GC analysis for the presence of methyl esters of 2-methylpentadecanoic acid failed to reveal a peak at the corresponding position, indicating that the conversion of the aldehyde to a fatty acid is performed by a separate enzyme. Hence, since the only activity of the puriﬁed enzyme is the speciﬁc cleavage of a carbon–carbon bond, we named it 2-hydroxyphytanoyl-CoA lyase, or HPCL. An apparent Km of 15 lM for 2-hydroxy-3-methylhexadecanoyl-CoA was calculated. When assayed in diﬀerent overlapping buﬀer systems, the lyase had a pH optimum between 7.5 and 8 in Tris buﬀer (1). Originally, HPCL had been puriﬁed in the absence of TPP, and the enzyme lost virtually all of its activity during puriﬁcation. The amino acid sequences of tryptic peptides from the puriﬁed and barely active HPCL suggested that the cleavage enzyme is related to a putative C. elegans protein, which displays homology to bacterial oxalyl-CoA decarboxylases. These enzymes, which have hitherto been described only in bacteria, catalyze the thiamine pyrophosphate (TPP)–dependent decarboxylation of oxalyl-CoA to formyl-CoA and CO2 (25,26). This homology suggested that HPCL might also require TPP, an as-yet-unrecognized cofactor for a-oxidation. In the presence of 0.8 mM Mg2+, optimum activity for the puriﬁed enzyme was reached at 20 lM TPP. Only minor stimulation by TPP was observed in a fresh liver homogenate (1.3-fold), and gradually more important stimulation of the lyase activity was observed as the enzyme fraction became more puriﬁed. In accordance with these ﬁndings, 20 lM TPP and 0.8 mM MgCl2 were from then on routinely included in the assay mixtures. The fact that enzyme activity decreased dramatically during puriﬁcation and could be restored only by adding TPP/Mg2+ might have been due to dissociation of monomers and concomitant loss of cofactors and points to a possible role of TPP in oligomerization of these proteins. Sequences derived from tryptic peptides of the puriﬁed rat protein were used as queries to recover human expressed sequence tags from the databases. The composite cDNA sequence of the human lyase contained an open reading frame of 1734 nucleotides encoding a polypeptide with a calculated molecular mass of 63,732 Da. Similarly to the bacterial oxalyl-CoA decarboxylases, a TPP-binding consensus domain could be identiﬁed in the C-terminal part of the lyase. The corresponding peptide sequences of this domain in the human, mouse, and rat enzyme, conform exactly with the TPP consensus domain of pyruvate decarboxylase of S. cerevisiae, acetolactate synthase of E. coli and of the putative oxalyl-CoA decarboxylases of O. formigenes, C. elegans, and S. cerevisiae (Fig. 3). Recombinant human protein, expressed in mammalian cells or in a yeast system, clearly exhibited lyase activity, whereas expression in a bacterial system did not result in a functionally active enzyme.

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Figure 3 Alignment of the cofactor-binding consensus domain in several TPPdependent enzymes. The corresponding peptide sequences of this domain in the human, rat, and mouse enzyme conform exactly with the TPP consensus domain of pyruvate decarboxylase of S. cerevisiae, acetolactate synthase of E. coli and of the putative oxalyl-CoA decarboxylase of O. formigenes.

Study of the substrate speciﬁcity of the His-fused recombinant human lyase revealed that the recombinant enzyme was active on 2-hydroxy3-methylhexadecanoyl-CoA (the analog of 2-hydroxyphytanoyl-CoA), and had much lower activity on 2-hydroxyhexadecanoyl-CoA. This last substrate, however, produced a very strong inhibition on the cleavage of 2-hydroxy-3-methylhexadecanoyl-CoA, most probably due to competition (27). No detectible activity was seen with 2-hydroxy-3-methylhexadecanoic acid, 3-methyl-hexadecanoic acid, or 3-methylhexadecanoyl-CoA, indicating that both a 2-hydroxygroup and a CoA moiety are necessary for lyase activity (27). At ﬁrst glance, the Hs HPCL sequence did not contain a C-terminal or N-terminal peroxisome-targeting signal (PTS). Because the C. elegans ortholog ends in a putative PTS1 (SKM) and since PRL, the C-terminal tripeptide of the S. cerevisiae ortholog, has been shown to bind to the human PTS1 import receptor (28), the C-terminal sequence SNM, which is also conserved in the mouse counterpart, could have a targeting function. Transfection studies with constructs coding for HPCL fused to GFP revealed that the ﬂuorescence localized to peroxisomes in ﬁbroblasts from PEX5+/ mice and to the cytosol in ﬁbroblasts from PEX5/ mice (29). The latter mice lack the PTS1 receptor (Pex5p) and do not import PTS1-containing proteins into their peroxisomes. Since a GFP construct containing only the last ﬁve amino acids of HPCL localized to peroxisomes in ﬁbroblasts from normal mice, we can conclude that targeting information is present within this pentapeptide and that SNM, preceded by a positive charge, is a hitherto-unrecognized PTS1. In the thiamine-dependent reaction catalyzed by HPCL, the formation of a carbanion, which reacts with carbon 1 of the substrate, is probably involved (Fig. 4). Ultimately this leads to the formation of formyl-CoA and a 2-methyl-branched fatty aldehyde. This mechanism makes HPCL the only

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Figure 4 In the thiamine-dependent reaction catalyzed by HPCL, the formation of a carbanion is supposed, which would react with carbon 1 of the substrate. Ultimately this leads to the formation of formyl-CoA and a 2-methyl-branched fatty aldehyde.

thiamine-dependent enzyme so far that has a 2-hydroxy compound instead of a 2-keto compound as substrate. Jones et al. (30) proposed an alternative mechanism for the conversion of 2-hydroxyphytanoyl-CoA, involving hydrolysis of the CoA-ester (peroxisomal thioesterases have been described) and a subsequent oxidation generating 2-ketophytanic acid, which would then be cleaved by HPCL, the enzyme we described. This hypothesis would turn HPCL into a more usual

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TPP-dependent enzyme because its substrate would then be a 2-keto compound. However, the activity of the required thioesterases and the 2-hydroxyacid oxidases on the substrates involved has never been demonstrated, whereas the activity of HPCL versus 2-hydroxy-3-methylacyl-CoA has unequivocally been proven. Moreover, if, according to the hypothesis of Jones et al. a 2-hydroxyacid oxidase were involved, no formyl-CoA/formate would be produced. This would be in contrast with the solid ﬁndings of several authors (6,10,11,14,22,31). The human HPCL gene has been mapped to chromosome 3p25 (Foulon, Vermeesch, Mannaerts, Casteels, and Van Veldhoven, unpublished results). The complete Hs HPCL gene spans 40.8 kb and contains 17 exons, with intron sizes ranging from 190 bp to 4700 bp. All exon–intron boundaries conform to the consensus rules, ending in an AG doublet and starting with a GT pair (32). Although several diseases are known to be associated with 3p25, none of these appear to be linked to HPCL. Moreover, up to now no patients with a deﬁcient HPCL have been identiﬁed. The mapping of the HPCL gene is a ﬁrst step toward the ﬁnding and diagnosis of such patients. As indicated before, Refsum disease can be attributed to several genetic defects (18), and the 3p25 locus might be another locus linked to this clinical picture. V. CONCLUSIONS AND FUTURE PERSPECTIVES 1. The fact that HPCL is clearly dependent on thiamine makes it plausible that the thiamine status of the cell will inﬂuence the aoxidation process. Preliminary experiments with cultured C6-glia cells or control human ﬁbroblasts under thiamine-deﬁcient conditions (generated either by the addition of 1 mM oxythiamine to the growth medium or by culturing cells in thiamine-depleted medium) showed a decrease of the overall ﬂux through the alpha-oxidation pathway (Foulon, Casteels, Mannaerts, Van Veldhoven, unpublished results). Would overall a-oxidation be deﬁcient in patients with thiamine deﬁciency as for example, in thiamine-responsive megaloblastic anemia (TRMA)? Would this lead to an accumulation of phytanic acid in these patients? 2. The fact that HPCL acts on a 2-hydroxy intermediate and not on a 2-keto compound diﬀerentiates it from the other thiamine-dependent enzymes and may shed new light on the reaction mechanism of this class of enzymes. 3. The reaction catalyzed by 2-hydroxyphytanoyl-CoA lyase is the ﬁrst example of a thiamine pyrophosphate–dependent reaction in peroxisomes. The mechanism for transport of thiamine or thiamine pyrophosphate into peroxisomes remains to be explored.

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ACKNOWLEDGMENTS This work was supported by grants from the Geconcerteerde onderzoeksacties van de Vlaamse Gemeenschap (GOA 94/98-12 and GOA 99/03-09) and from the Fonds voor Wetenschappelijk Onderzoek-Vlaanderen (G-0239.98 and G.0115.02). V.F. was supported by a fellowship from the Fonds voor Wetenschappelijk Onderzoek-Vlaanderen. Mr. Luc Govaert, Ms. Wendy Geens, and Mr. Stanny Asselberghs are gratefully acknowledged for their technical support. ABBREVIATIONS PAHX HPCL ARD PTS TPP

phytanoyl-CoA hydroxylase 2-hydroxyphytanoyl-CoA lyase adult Refsum disease peroxisome targeting signal thiamine pyrophosphate

REFERENCES 1.

2.

3.

4. 5. 6.

7.

V Foulon, VD Antonenkov, K Croes, E Waelkens, GP Mannaerts, PP Van Veldhoven, M Casteels. Puriﬁcation, molecular cloning, and expression of 2hydroxyphytanoyl-CoA lyase, a peroxisomal thiamine pyrophosphate–dependent enzyme that catalyzes the carbon–carbon bond cleavage during alphaoxidation of 3-methyl-branched fatty acids. Proc Natl Acad Sci USA 96:10039– 10044, 1999. E Klenk, W Kahlke. U¨ber das Vorkommen der 3, 7, 11, 15-Tetramethylhexadecansa¨ure (Phytansa¨ure) in den Cholesterinestern und andere Lipidfractionen der Organe bei einem Krankheitsfall unbekannter Genese (Verdacht auf Heredopathia Atactica Polyneuritiformis [Refsum Syndrome]). Hoppe Zeilers Z Physiol Chem 333:133–139, 1963. D Steinberg. Refsum disease. In: CR Scriver, AL Beudet, WS Sly, D Valle, eds. The Metabolic and Molecular Bases of Inherited Disease. 7th ed. New York: McGraw-Hill, 1995, pp 2361–2369. GP Mannaerts, PP Van Veldhoven, M Casteels. Peroxisomal lipid degradation via a- and h-oxidation in mammals. Cell Biochem Biophys 32:73–87, 2000. SR Terlecky, M Fransen. How peroxisomes arise. Traﬃc 1:465–473, 2000. A Poulos, P Sharp, H Singh, DW Johnson, WF Carey, C Easton. Formic acid is a product of the a-oxidation of fatty acids by human skin ﬁbroblasts: deﬁciency of formic acid production in peroxisome-deﬁcient ﬁbroblasts. Biochem J 292:457– 461, 1993. PA Watkins, AE Howard, SJ Gould, J Avigan, SJ Mihalik. Phytanic acid activation in rat liver peroxisomes is catalyzed by long-chain acyl-CoA synthetase. J Lipid Res 37:2288–2295, 1996.

536 8.

9.

10.

11.

12.

13. 14.

15.

16.

17.

18.

19.

20.

21.

Casteels et al. K Pahan, J Cofer, P Baliga, I Singh. Identiﬁcation of phytanoyl-CoA ligase as a distinct acyl-CoA ligase in peroxisomes from cultured human skin ﬁbroblasts. FEBS Lett 322:101–104, 1993. SJ Steinberg, SJ Wang, DG Kim, SJ Mihalik, PA Watkins. Human very-longchain acyl-CoA synthetase: cloning, topography, and relevance to branchedchain fatty acid metabolism. Biochem Biophys Res Commun 257:615–621, 1999. SJ Mihalik, AM Rainville, PA Watkins. Phytanic acid a-oxidation in rat liver peroxisomes. Production of a-hydroxyphytanoyl-CoA and formate is enhanced by dioxygenase cofactors. Eur J Biochem 232:545–551, 1995. K Croes, M Casteels, GP Mannaerts, PP Van Veldhoven. a-Oxidation of 3methyl-substituted fatty acids in rat liver. Production of formic acid instead of CO2, cofactor requirements, subcellular localization and formation of a 2hydroxy-3-methylacyl-CoA intermediate. Eur J Biochem 240:674–683, 1996. GA Jansen, SJ Mihalik, PA Watkins, HW Moser, C Jakobs, S Denis, RJA Wanders. Phytanoyl-CoA hydroxylase is present in human liver, located in peroxisomes, and deﬁcient in Zellweger syndrome: direct, unequivocal evidence for the new, revised pathway of phytanic acid a-oxidation in humans. Biochem Biophys Res Commun 229:205–210, 1996. SJ Mihalik, JC Morrell, D Kim, KA Sacksteder, PA Watkins, SJ Gould. Identiﬁcation of PAHX, a Refsum disease gene. Nat Genet 17:185–189, 1997. K Croes, PP Van Veldhoven, GP Mannaerts, M Casteels. Production of formyl-CoA during peroxisomal a-oxidation of 3-methyl-branched fatty acids. FEBS-Letters 407:197–200, 1997. K Croes, M Casteels, S Asselberghs, P Herdewijn, GP Mannaerts, PP Van Veldhoven. Formation of a 2-methyl-branched fatty aldehyde during peroxisomal a-oxidation. FEBS Lett 412:643–645, 1997. NM Verhoeven, DSM Schor, HJ ten Brink, RJA Wanders, C Jakobs. Resolution of the phytanic acid a-oxidation pathway: identiﬁcation of pristanal as product of the decarboxylation of 2-hydroxyphytanoyl-CoA. Biochem Biophys Res Commun 237:33–36, 1997. AS Wierzbicki, MD Loyd, CJ Schoﬁeld, MD Feher, FB Gibberd. Refsum’s disease: a peroxisomal disorder aﬀecting phytanic acid a-oxidation. J Neurochem 80:727–735, 2002. AS Wierzbicki, J Mitchell, M Lambert-Hammill, M Hancock, J Greenwood, MC Sidey, J de Belleroche, FB Gibberd. Identiﬁcation of genetic heterogeneity in Refsum’s disease. Eur J Hum Genet 8:649–651, 2000. GA Jansen, R Ofman, S Ferdinandusse, L Ijlst, AO Muijsers, OH Skjeldal, O Stokke, C Jakobs, GTN Besley, JE Wraith, RJA Wanders. Refsum disease is caused by mutations in the phytanoyl-CoA hydroxylase gene. Nat Genet 17:190–193, 1997. AS Wierzbicki, M Lambert-Hammill, J Mitchell, M Sidey, MD Feher, J de Belleroche, FB Gibberd. Identiﬁcation of a second locus for Refsum’s disease at chromosome 6p22-24. Am J Human Genetics 67(suppl 2):1644, 2000. S Ferdinandusse, S Denis, PT Clayton, A Graham, JE Rees, JT Allen, BN McLean, AY Brown, P Vreken, HR Waterham, RJA Wanders. Mutations in

Thiamine Pyrophosphate in the Metabolism of Fatty Acids

22.

23.

24.

25. 26.

27.

28.

29.

30.

31.

32.

537

the gene encoding peroxisomal alpha-methylacyl-CoA racemase cause adultonset sensory motor neuropathy. Nat Genet 24:188–191, 2000. M Casteels, K Croes, PP Van Veldhoven, GP Mannaerts. Aminotriazole is a potent inhibitor of a-oxidation of 3-methyl-substituted fatty acids in rat liver. Biochem Pharmacol 48:1973–1975, 1994. PP Van Veldhoven, S Huang, HJ Eyssen, GP Mannaerts. The deﬁcient degradation of synthetic 2- and 3-methyl-branched fatty acids in ﬁbroblasts from patients with peroxisomal disorders. J Inher Metab Dis 16:381–391, 1993. VD Antonenkov, SJ Pirozhkov, LF Panchenko. Intraparticulate localization and some properties of a cloﬁbrate-induced peroxisomal aldehyde dehydrogenase from rat liver. Eur J Biochem 149:159–167, 1985. AL Baetz, MJ Allison. Puriﬁcation and characterization of oxalyl-coenzyme A decarboxylase from Oxalobacter formigenes. J Bacteriol 171:2605–2608, 1989. HY Lung, AL Baetz, AB Peck. Molecular cloning, DNA sequence, and gene expression of the oxalyl-coenzyme A decarboxylase gene, oxc, from the bacterium Oxalobacter formigenes. J Bacteriol 176:2468–2472, 1994. V Foulon. a-Oxidation of 3-methyl-branched fatty acids: study of the enzymes involved in the reaction sequence. PhD dissertation. Acta Biomedica Lovaniensa. Leuven University Press, 2001. G Lametschwandtner, C Brocard, M Fransen, PP Van Veldhoven, J Berger, A Hartig. The diﬀerence in recognition of terminal tripeptides as peroxisomal targeting signal 1 between yeast and human is due to diﬀerent aﬃnities of their receptor Pex5p to the cognate signal and to residues adjacent to it. J Biol Chem 273:33635–33643, 1998. M Baes, P Gressens, E Baumgart, P Carmeliet, M Casteels, M Fransen, P Evrard, D Fahimi, PE Declercq, D Collen, PP Van Veldhoven, GP Mannaerts. A mouse model for Zellweger syndrome. Nature Genetics 17:49–56, 1997. JM Jones, JC Morell, SJ Gould. Identiﬁcation and characterization of HAOX1, HAOX2, and HAOX3, three human peroxisomal 2-hydroxy acid oxidases. J Biol Chem 275:12590–12597, 2000. NM Verhoeven, DSM Schor, SF Previs, H Brunengraber, C Jakobs. Stable isotope studies of phytanic acid a-oxidation: in vivo production of formic acid. Eur J Pediatr 156:S83–S87, 1997. SM Mount. A catalogue of splice junction sequences. Nucleic Acid Res 10:459– 472, 1982.

30 Pathogenesis of Selective Neuronal Loss in Wernicke–Korsakoff Syndrome: Role of Oxidative Stress Paul Desjardins and Roger F. Butterworth University of Montreal, Montreal, Quebec, Canada

I. INTRODUCTION Wernicke–Korsakoﬀ syndrome (WKS) is a neuropsychiatric disorder characterized by ophthalmoplegia (horizontal gaze palsy, ptosis), gait ataxia, and a global confusional state. WKS is a common complication of chronic alcoholism and is also encountered in patients with HIV-AIDS, gastric carcinoma, and other disorders associated with grossly impaired nutritional status. There is abundant evidence to suggest that WKS results from thiamine deﬁciency. In chronic alcoholism, thiamine deﬁciency results from poor diet, impaired absorption of thiamine from the gastrointestinal tract, and a loss of liver thiamine stores resulting from alcoholic liver disease. In addition, alcohol may impair the phosphorylation of thiamine both in peripheral tissues and in brain. Neuropathologic evaluation of brain tissue from WKS patients reveals a pattern of selective damage to mammillary bodies, thalamus, and pons (1). Cellular changes include neuronal loss, astrocytic proliferation, and microglial activation. The cause of this distinctive pattern of neurodegeneration is unknown, but several theories involving cellular energy failure, lactate 539

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accumulation, blood–brain barrier breakdown, and exitotoxicity have been proposed. In experimental animals exposed to thiamine deﬁciency, both apoptotic and necrotic patterns of neuronal cell death have been reported. This chapter reviews evidence that oxidative stress is a major pathophysiologic phenomenon implicated in neurodegeneration in WKS and that the mechanisms involved are common to many neurodegenerative CNS disorders. II. THIAMINE ESTERS AND THIAMINE-DEPENDENT ENZYMES In the 1930s, Peters and coworkers showed that thiamine deﬁciency in pigeons resulted in the accumulation of lactate in the brainstem (2), a region of brain known to be selectively lesioned due to thiamine deﬁciency in this species. Furthermore, he showed that the addition of small quantities of crystalline thiamine to the excised brain tissue from thiamine-deﬁcient birds in vitro resulted in correction of the metabolic defect and in normalization of lactate levels. These ﬁndings led Peters to formulate the concept of ‘‘the biochemical lesion’’ in thiamine deﬁciency. Later studies showed that the enzyme defect responsible for the ‘‘biochemical lesion’’ was a-ketoglutarate dehydrogenase complex (a-KGDHC) rather than pyruvate dehydrogenase complex (PDHC) (as had initially been suspected). a-KGDHC, PDHC, and transketolase are major thiamine diphosphate (TDP)–dependent enzymes involved in glucose metabolism (Fig. 1). In brain, thiamine occurs predominantly (over 80%) in the form of TDP, the remainder being made up of thiamine monophosphate (10%) and thiamine triphosphate (5–10%), with only trace amounts of free thiamine. Thiamine is transported into brain and phosphorylated by the action of thiamine pyrophosphokinase, and inhibition of this enzyme by the thiamine antagonist pyrithiamine or by ethanol results in decrease synthesis of TDP and, consequently, decreased activities of thiamine-dependent enzymes. Treatment of experimental animals with pyrithiamine results in a generalized reduction of TDP concentrations throughout brain (3) but in an early selective loss in activity of a a-KGDHC. Decreased activities of a-KGDHC following treatment with pyrithiamine led to decreased synthesis of glucosederived excitatory and inhibitory amino acids in brain, including glutamate, aspartate, and GABA, and a concomitant increase in lactate and alanine (4), all of which are consistent with decreased ﬂux of carbon through the tricarboxylic acid cycle (Fig. 1). Both the a-KGDHC decreases and the changes in synthesis of amino acids are reversible following thiamine rehabilitation in pyrithiamine-treated animals (4), suggesting that these changes are an integral part of ‘‘the biochemical lesion’’ in thiamine deﬁciency. In a study of thiamine-dependent enzymes in postmortem brain tissue from alcoholic patients, it was reported that decreased activities of PDHC,

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Figure 1 Decreased activities of a-KGDHC due to thiamine deﬁciency results in decreased synthesis of glutamate, aspartate, and GABA, in mitochondrial uncoupling, and in a cellular energy deﬁcit. PDHC: pyruvate dehydrogenase complex; TK: transketolase.

a-KGDHC, and transketolase were conﬁned to those cases in with a neuropathologic diagnosis of WKS had been made (5). Alcoholic patients without WKS manifested brain thiamine-dependent enzyme activities within normal limits. Furthermore, activities of a non-thiamine-dependent enzyme glutamate dehydrogenase were found to be unchanged in both WKS and nonWKS alcoholic brain. These ﬁndings provide evidence, for the ﬁrst time, that reduction in thiamine-dependent enzymes is implicated in the pathogenesis of the selective neuronal cell loss that is characteristic of WKS in humans.

III. MITOCHONDRIAL DYSFUNCTION IN THIAMINE DEFICIENCY Consistent with reductions in activity of a-KGDHC, oxidative decarboxylation of a-ketoglutarate (and pyruvate) are reportedly decreased in isolated

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mitochondria from the brains of pyrithiamine-treated rats (6), and, as was the case in the in vitro studies, normal rates of decarboxylation and enzyme activities were promptly restored after the addition of TDP to the mitochondrial preparations. Subsequent studies in mitochondrial preparations from thiamine-deﬁcient animals showed that state 3 respiration was decreased using a-ketoglutatate as substrate but was unchanged using succinate (6) (which is oxidized independent of TDP-dependent enzymes; see Fig. 1) Several studies have been conducted to elucidate the cellular metabolic consequences of thiamine deﬁciency in relation to mitochondrial function using cultured neuronal cell preparations. Cultured cerebellar granule cells exposed to pyrithiamine manifest signiﬁcant reductions in TDP accompanied by reduced activities of a-KGDHC, increased lactate production indicative of decreased tricarboxylic acid cycle ﬂux, and a lowering of cellular pH (7). In a study in which cultures of neuroblastoma cells were exposed to the thiamine transport inhibitor amprolium, decreases in activity of a-KGDHC led to decreased oxygen consumption, uncoupling of mitochondria, and disorganization of cristae, all of which could be restored following the addition of thiamine or succinate (8). More prolonged exposure of either neuroblastoma or primary cultures of cerebellar granule cells to thiamine deﬁciency, however, led to depolarization, lactate accumulation, decreased ATP production, and cell necrosis. Studies in vivo of pyrithiamine-treated rats likewise revealed disintegration of mitochondria and chromatin clumping in degenerating diencephalic neurons (9). Decreases of ATP in brainstem have also been described in pyrithiamine-treated animals (10). As a result of these studies, impairment of cerebral energy metabolism leading to brain lactate accumulation and acidosis have been proposed to explain the phenomenon of selective neuronal cell loss due to thiamine deﬁciency. Indeed, signiﬁcant acidosis has been described in vulnerable brain structures such as mammillary bodies, thalamus, and pons of pyrithiamine-treated rats (11).

IV. NMDA RECEPTOR-MEDIATED EXITOTOXICITY IN THIAMINE DEFICIENCY The neuropathologic lesions observed in severe thiamine deﬁciency resemble to some extent those observed in conditions of anoxia-ischemia as well as in NMDA receptor-mediated glutamate-induced excitotoxicity. In favor of such a mechanism in thiamine deﬁciency, increased concentrations of glutamate have been reported in brain extracellular (dialysate) ﬂuid from pyrithiaminetreated rats (12) and, more recently, decreases in expression of astrocytic glutamate transporters was reported in the brains of these animals (13), a phenomenon that could explain the increased extracellular brain glutamate

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concentrations. Also consistent with an NMDA receptor-mediated excitotoxic mechanism in the pathogenesis of the selective neuronal cell loss due to thiamine deﬁciency were the ﬁndings that administration of MK801, an NMDA receptor antagonist, resulted in attenuation of necrotic damage in thalamus of pyrithiamine-treated animals (14). However, a subsequent study attributed at least some of the neuroprotective eﬀects of MK801 to its anticonvulsant and hypothermic properties (15).

V. OXIDATIVE STRESS: ROLE IN THIAMINE-DEFICIENCYRELATED NEURONAL CELL DEATH There is an increasing body of evidence to suggest that oxidative stress contributes to the cellular energy deﬁcit and cell death in thiamine deﬁciency. Increased production of oxygen free radicals (reactive oxygen species) have been reported in brain in pyrithiamine-treated rats (16), and oxidative stress is related to the pathology in human WKS characterized by increased neuronal peroxidase activity. Changes that are consistent with oxidative stress in thiamine deﬁciency are the ﬁndings of increased amounts of hemoxygenase and ICAM-1 (17) as well as the ﬁnding of microglial activation (18). Moreover, selective induction of the endothelial isoform of nitric oxide synthase (eNOS) has been reported in the brains of pyrithiamine-treated animals (Fig. 2). Increases of eNOS expression were particularly apparent in medial thalamus of thiamine-deﬁcient animals; no signiﬁcant changes were observed in cerebral cortex. These ﬁndings suggest that an early, region-selective insult in thiamine deﬁciency is increased production of nitric oxide (NO) by vascular endothelial cells. Indeed the vascular endothelium has been proposed as major site of free-radical production in both experimental thiamine deﬁciency and in WKS in humans. Using an immunohistochemical approach, an increase of eNOS immunostaining was observed in thalamic microvessels of pyrithiamine-treated mice (19). Increased iNOS immunostaining of microglia was also reported in the brains of these animals. These ﬁndings of increased NOS expression and NO production are likely to be of pathophysiologic signiﬁcance since NO can react with the superoxide radical (O2) to form peroxynitrite (ONOO), which is directly toxic to neurons. Its toxicity results from its ability to nitrate tyrosine residues and to cross-link thiol groups in proteins. Increased nitrotyrosine immunolabeling has been described in neuronal processes in thalamus of pyrithiamine-treated animals (20), consistent with its role in the pathogenesis of selective neuronal cell death due to thiamine deﬁciency. Furthermore, the targeted disruption of the eNOS gene was shown to result in a reduction in the extent of the necrotic lesions in thalamus of pyrithiamine-treated animals (19).

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Figure 2 Increased expression of endothelial nitric oxide synthase (eNOS) mRNA in the medial thalamus of thiamine-deﬁcient rats and pair-fed controls. Total RNA was extracted from the medial thalamus of thiamine-deﬁcient rats (TD) or from pair-fed controls. h-Actin (347 bp) and eNOS (351 bp) were reverse-transcribed and ampliﬁed by PCR for 18 and 32 cycles, respectively. Lane 1: molecular weight standard; lanes 3–6: pair-fed controls; lanes 8–15: symptomatic TD rats; lanes 2 and 7: AMV reverse transcriptase was omitted from the reaction mixture as a negative control.

VI. OXIDATIVE STRESS AND MITOCHONDRIAL DYSFUNCTION: VISCIOUS CYCLE INVOLVED IN NEURODEGENERATION DUE TO THIAMINE DEFICIENCY Increases in NO derived from eNOS induction have been shown to result in decreased activities of mitochondrial enzymes, including a-KGDHC. For example, an inhibitory eﬀect of NO on cytochrome c oxidase activities has

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been reported in both isolated mitochondria and various neuronal preparations (21). Base upon these observations, it is likely that NO and peroxynitrite-mediated mitochondrial dysfunction and damage play a role in neuronal cell loss. Evidence in favor of such a mechanism is compelling. For example, a-KGDHC is sensitive to oxidation products that occur in Parkinson’s disease (17), and thiamine-dependent enzymes, including a-KGDHC are reduced in activity in autopsied brain tissue from patients with Alzheimer’s disease. NO exerts an inhibitory eﬀect on cytochrome c oxidase and aKGDHC activities in microglial cells, and both microglial activation and reductions in a-KGDHC have consistently been described in a wide range of neurodegenerative disorders, including Alzheimer’s disease (17), ischemic brain injury (22), and WKS (5). In the case of WKS, these mechanisms may lead to a viscious cycle, whereby thiamine deﬁciency results in decreased availability of cofactor (TDP), leading to reduction in activity of a-KGDHC, a rate-limiting tricarboxylic acid cycle enzyme that in turn results in mitochondrial uncoupling. Simultaneously, NO produced by activation of endothelial and microglial NOS combines with the superoxide radical (O 2 ) to form peroxynitrite, both

Figure 3 Cellular energy failure and possible mechanisms whereby eNOS induction could participate in the pathogenesis of selective neuronal cell loss due to thiamine deﬁciency.

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of which result in mitochondrial dysfunction and further reductions in activities of a-KGDHC (Fig. 3). Further evidence in support of a role of oxidative stress in the pathogenesis of selective neuronal loss in thiamine deﬁciency is provided by the report of a neuroprotective eﬀect of L-deprenyl (23), an agent with potent oxygen free radical scavenging properties. Interestingly, L-deprenyl has been shown to cause a slowing in the evolution of the neurological defects in Parkinson’s disease (24) and in Alzheimer’s disease (25), two neurodegenerative disorders associated with decreased activities of a-KGDHC. REFERENCES 1.

CG Harper, RF Butterworth. Nutritional and Metabolic Disorders. In: DI Graham, PL Lantos, eds. Greenﬁeld’s Neuropathology. London: Arnold, 1997, pp. 601–655. 2. RA Peters. The biochemical lesion in vitamin B1 deﬁciency. Application of modern biochemical analysis in its diagnosis. Lancet 1:1161–1164, 1936. 3. M He´roux, RF Butterworth. Regional alterations of thiamine phosphate esters and of thiamine diphosphate–dependent enzymes in relation to function in experimental Wernicke’s encephalopathy. Neurochem Res 20:87–93, 1995. 4. RF Butterworth, M He´roux. Eﬀect of pyrithiamine treatment and subsequent thiamine rehabilitation on regional cerebral amino acids and thiamine-dependent enzymes. J Neurochem 52:1079–1084, 1989. 5. RF Butterworth, JJ Kril, C Harper. Thiamine-dependent enzyme changes in brain of alcoholics: relationship to Wernicke–Korsakoﬀ syndrome. Alcohol Clin Exp Res 17:1084–1088, 1993. 6. WD Jr. Parker, R Haas, DA Stumpf, J Parks, LA Eguren, C Jackson. Brain mitochondrial metabolism in experimental thiamine deﬁciency. Neurology 34:1477–1481, 1984. 7. P Pannunzio, AS Hazell, M Pannunzio, KV Rama Rao, RF Butterworth. Thiamine deﬁciency results in metabolic acidosis and energy failure in cerebellar granule cells: an in vitro model for the study of cell death mechanisms in Wernicke’s encephalopathy. J Neurosci Res 62:286–292, 2000. 8. L Bettendorﬀ, F Sluse, G Goessens, P Wins, T Grisar. Thiamine deﬁciency– induced partial necrosis and mitochondrial uncoupling in neuroblastoma cells are rapidly reversed by addition of thiamine. J Neurochem 65:2178–2184, 1995. 9. SX Zhang, GS Weilersbacher, SW Henderson, T Corso, JW Olney, PJ Langlais. Excitotoxic cytopathology, progression, and reversibility of thiamine deﬁciency– induced diencephalic lesions. J Neuropathol Exp Neurol 54:255–267, 1995. 10. H Aikawa, IS Watanabe, T Furuse, Y Iwasaki, E Satoyoshi, T Sumi, T Moroji. Low energy level in thiamine-deﬁcient encephalopathy. J Neuropathol Exp Neurol 43:276–287, 1984. 11. AM Hakim. The induction and reversibility of cerebral acidosis in thiamine deﬁciency. Ann Neurol 16:673–679, 1984.

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12. AS Hazell, RF Butterworth, AM Hakim. Cerebral vulnerability is associated with selective increase in extracellular glutamate concentration in experimental thiamine deﬁciency. J Neurochem 61:1155–1158, 1993. 13. AS Hazell, KV Rao, NC Danbolt, DV Pow, RF Butterworth. Selective downregulation of the astrocyte glutamate transporters GLT-1 and GLAST within the medial thalamus in experimental Wernicke’s encephalopathy. J Neurochem 78:560–568, 2001. 14. PJ Langlais, RG Mair. Protective eﬀects of the glutamate antagonist MK-801 on pyrithiamine-induced lesions and amino acid changes in rat brain. J Neurosci 10:1664–1674, 1990. 15. KG Todd, RF Butterworth. Evaluation of the role of NMDA-mediated exitotoxicity in the selective neuronal loss in experimental Wernicke encephalopathy. Exp Neurol 149:130–138, 1997. 16. PJ Langlais, G Anderson, SX Guo, SC Bondy. Increased cerebral free radical production during thiamine deﬁciency. Metab Brain Dis 12:137–143, 1997. 17. GE Gibson, H Zhang. Interactions of oxidative stress with thiamine homeostasis promote neurodegeneration. Neurochem Int 40:493–504, 2002. 18. KG Todd, RF Butterworth. Early microglial response in experimental thiamine deﬁciency: an immunohistochemical analysis. Glia 25:190–198, 1999. 19. NY Calingasan, GE Gibson. Vascular endothelium is a site of free radical production and inﬂammation in areas of neuronal loss in thiamine-deﬁcient brain. Ann NY Acad Sci 903:353–356, 2000. 20. NY Calingasan, LC Park, LL Calo, RR Triﬁletti, SE Grandy, GE Gibson. Induction of nitric oxide synthase and microglial responses precede selective cell death induced by chronic impairment of oxidative metabolism. Am J Pathol 153:599–610, 1998. 21. GC Brown. Regulation of mitochondrial respiration by nitric oxide inhibition of cytochrome C oxidase. Biochim Biophys Acta 1504:46–57, 2001. 22. W Cao, JM Carney, A Duchon, RA Floyd, M Chevion. Oxygen free radical involvement in ischemia and reperfusion injury to brain. Neurosci Lett 88:23– 238, 1988. 23. KG Todd, RF Butterworth. Increased neuronal cell survival after l-deprenyl treatment in experimental thiamine deﬁciency. J Neurosci Res 52:240–246, 1998. 24. The Parkinson Group Study. Eﬀects of tocopherol and deprenyl on the progression of disability in early Parkinson’s disease. New Engl J Med 328:176–183, 1993. 25. LS Schneider, VE Pollock, MF Zemansky, RP Gleason, R Palmer, RB Sloane. A pilot study of low-dose l-deprenyl in Alzheimer’s disease. J Geriatr Psychiatry Neurol 4:143–148, 1991.

31 Thiamine-Responsive Megaloblastic Anemia Syndrome: Clinical Aspects and Molecular Genetics Kimihiko Oishi, George A. Diaz, and Bruce D. Gelb Mount Sinai School of Medicine, New York, New York, U.S.A.

I.

INTRODUCTION

Rogers et al. described the ﬁrst thiamine-responsive megaloblastic anemia syndrome (OMIM 249270) patient in 1969. The patient was an 11-year-old girl who presented with megaloblastic anemia, diabetes mellitus, amino aciduria, and sensorineural deafness without clear cause (1). The administration of various vitamins and cofactors revealed that this disorder responded only to treatment with thiamine. Viana and Carvalho reported a similar case of this combination of symptoms in 1978 (2). Their patient was a 6-year-old female with anemia responsive to pharmacological doses of oral thiamine. She presented with latent diabetes mellitus, sensorineural deafness, and situs inversus totalis. Her parents were ﬁrst cousins, suggesting autosomal recessive inheritance. With the ascertaining of several additional cases, it was noted that consanguineous mating was a common factor, leading to the acceptance of this mode of inheritance. Because one of the main features of this disorder, megaloblastic anemia, was responsive to oral thiamine, it was named thiamine-responsive megaloblastic anemia syndrome, or TRMA. It has also been referred to eponymically as Rogers syndrome. 549

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Biochemical studies revealed that TRMA erythrocytes and ﬁbroblasts lacked the high-aﬃnity saturable component of thiamine uptake (3–5), leading to the proposal that the primary defect in TRMA was an absence of a thiamine transporter. Since no mammalian thiamine transporters had been identiﬁed, the research groups of Neufeld and Cohen adopted a positional cloning strategy. They performed a genome scan using several kindreds with TRMA, which successfully mapped the trait to 1q23.2–23.3 (6). After those researchers and our group reﬁned the TRMA critical region to a more tractable size (7,8), the three groups independently identiﬁed the disease-causing gene, SLC19A2, using various positional cloning/candidacy strategies (9–11). SLC19A2 encodes the high-aﬃnity thiamine transporter, THT1 (10,12). The pathogenesis of TRMA remains somewhat unclear, leading our group to turn to animal modeling. We cloned the orthologous mouse gene, Slc19a2 (13), and then created a targeted disruption of this gene through homologous recombination in embronic stem cells (14). Characterization studies of mice homozygous for this knockout allele revealed that megaloblastosis, deafness, and diabetes mellitus occurred when the animals were maintained on a thiamine-depleted diet (14). These studies provided some insights into the pathogenesis of diabetes mellitus in TRMA, and future explorations are expected to provide additional information about all aspects of the phenotype. II.

CLINICAL MANIFESTATIONS

Megaloblastic anemia, sensorineural deafness, and diabetes mellitus are the main features of TRMA. In addition to this triad, thrombocytopenia, optic atrophy, congenital heart defects, cardiomyopathy, and acute ischemic stroke have been described in some patients (1,3,15–18). Development does not appear to be aﬀected. In general, patients present in the ﬁrst year of life with the anemia. The diabetes mellitus and deafness may be detected at presentation or can occur later in early childhood. Signs of beriberi, like edema and peripheral neuropathy, are not observed in these patients, which is notable, given the relationship of TRMA to thiamine transport. The hematological characteristics of TRMA are fairly uniform in the reported cases. Peripheral blood counts show persistently low hemoglobin levels and high mean corpuscular volume (MCV). There is no abnormality in leukocyte numbers or diﬀerential. Peripheral blood smears reveal macrocytosis, anisocytosis, poikilocytosis, and nuclear hypersegmentation of granulocytes. Plasma iron concentration and iron-binding capacity are within the normal range. This macrocytic anemia cannot be attributed to deﬁciency of folate or vitamin B12 because plasma levels of these vitamins have been normal in TRMA patients. Bone marrow aspirates show dysplastic hema-

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topoiesis with a reversed myeloid : erythroid (M : E) ratio due to erythroid hyperplasia. In the erythroid lineage, numerous megaloblasts are evident with dyssynchronous nuclear and cytoplasmic maturation, with a lag in the former. This constellation of ﬁndings is quite similar to those observed in folate and vitamin B12 deﬁciencies, which are attributable to impaired DNA synthesis in cells with rapid turnover. While megaloblastic bone marrow changes are an essential feature of TRMA, several variations have been reported. In some patients, ring sideroblasts are seen in the bone marrow (1,19). Bazarbachi et al. described two patients with tri-lineage myelodysplastic changes in a bone marrow aspirate and trephine biopsy (20). Diabetes mellitus typically presents by the fourth year of life in children with TRMA. The initial presentation is usually glycosuria and hyperglycemia. Diabetic ketoacidosis has not been reported, and the diagnosis of diabetes mellitus is often made incidentally, which is distinctly diﬀerent from typical type I disease. Fasting hyperglycemia is common, and glucose intolerance is observed on oral glucose tolerance test (16,21). In some cases, patients show latent diabetes mellitus with mild glucose intolerance (2). Initially, insulin secretion is retained, and plasma insulin measurements reveal normal values following glucose stimulation. The use of oral hypoglycemic agents to treat the hyperglycemia can be successful for several years after the onset of diabetes mellitus (1,22). In the long term, however, most patients become insulin dependent due to inadequate insulin secretion. In light of the absence of ketoacidosis without insulin therapy, normal insulin response to oral glucose loading, and initial responsiveness to oral hypoglycemic agents, the diabetes mellitus associated with TRMA has been attributed to peripheral insulin insensitivity similar to type II disease. The eventual insulin dependence suggests the exhaustion of insulin secretory capacity, perhaps due to destruction of h islet cells in the pancreas (22). Deafness is observed in all reported TRMA cases. Audiograms show that this aspect of the phenotype results from profound bilateral sensorineural hearing loss (2,20,21,23). Patients often need hearing aids and suﬀer from the expected diﬃculties in speech and language development. Thiamine therapy appears to stabilize the deafness in the short term, but reversibility has been documented only rarely. Long-term progression of hearing loss has not been investigated in TRMA, since nearly all patients are profoundly deaf at presentation. III. TREATMENT The mainstay of therapy for TRMA is the administration of pharmacological doses of oral thiamine. Typical dosing is 20–100 mg of thiamin-HCI or 50 mg of the lipophilic form of thiamine (benzoyl-oxymethyl-thiamine, BOM)

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orally. As described by Rogers et al. (1), oral thiamine is very eﬀective for treating the megaloblastic anemia. It increases the reticulocyte count, hemoglobin, and other peripheral hematological indices. The reticulocytic response to thiamine therapy becomes apparent after ﬁve to ten days, and the hemoglobin level returns to normal in 10–14 days. The MCV decreases with therapy, and erythrocyte morphology returns to almost normal, with only small abnormalities persisting. Since thiamine therapy does not alter the fundamental biological defect in TRMA, cessation of thiamine administration or compliance failure results in an immediate recurrence of the hematological abnormalities. High-dose thiamine treatment also has a favorable eﬀect on the diabetes mellitus. Initially, thiamine therapy normalizes blood glucose levels and glucose tolerance test curves (1,2,16,19,21,22). Unlike the hematologic manifestations, however, normalization of function is only temporary. Nonetheless, thiamine therapy may prolong the duration prior to the initiation of insulin therapy or, for patients already insulin dependent, reduce or abolish the insulin requirement (1,2,16,19,21). In two patients who were treated from early in life, metabolic control deteriorated around the time of puberty, resulting in the need for oral hypoglycemic agents and, eventually, insulin administration (22). Of the three principal aspects of the TRMA phenotype, the sensorineural deafness is ameliorated least well by thiamine therapy. In fact, there is no improvement in hearing function in most TRMA cases. The response to thiamine is usually limited to a slight improvement in the auditory-evoked response or an arrest of the hearing deterioration (21,23). This suggests that the pathology of the deafness is irreversible once present clinically (23). In 1983, Tenconi and colleagues identiﬁed a lipophilic form of thiamine, BOM, and Poggi’s group introduced it for treating TRMA (22,24). This compound was posited as a superior therapeutic modality because it crosses biological membranes more readily, allowing higher intracellular concentrations of thiamine. The eﬃcacy of BOM as therapy for TRMA was documented in several studies, although the diﬀerences when compared to standard thiamine-HC1 therapy were not dramatic (3,4,22,23). In one patient, BOM use resulted in an improvement of the auditory evoked potentials, a response that has never been observed with thiamine-HC1 (22). It remains to be determined whether initiation of thiamine therapy at birth will prevent deafness, although this measure is clearly indicated in cases diagnosed prenatally or in early infancy. IV.

BIOCHEMICAL ASPECTS

The deﬁcit of a high-aﬃnity thiamine transporter, resulting in low intracellular thiamine concentrations, is the main biochemical defect of TRMA.

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Rindi’s group ﬁrst showed that the levels of thiamine compounds (thiamine, thiamine monosphosphate, and thiamine pyrophosphate) were signiﬁcantly decreased in red blood cells while serum thiamin levels were normal (3,4,23). Thiamine uptake studies demonstrated that active transport of thiamine was absent in TRMA erythrocytes and ghosts. Neufeld’s group observed similar phenomena using skin ﬁbroblasts from TRMA patients (5). TRMA ﬁbroblasts lacked the high-aﬃnity uptake of thiamine and had signiﬁcantly increased apoptosis when cultured in a physiological thiamine concentration. A precise understanding of the pathogenesis of TRMA remains elusive. With respect to the anemia, deoxyuridine suppression testing indicated that the megaloblastic changes were not secondary to an impairment of the methylation of deoxyuridylate (19). Incorporation studies revealed no evidence of either DNA or RNA synthesis defects in the marrow cells. Based on these results, it was suggested that the megaloblastic changes were not caused by a primary defect in nucleic acid synthesis but were a poorly deﬁned secondary manifestation. In 2001, Neufeld and colleagues proposed that the abnormal erythropoiesis resulted from inadequate synthesis of ribose-5-phosphate, based on their studies with TRMA ﬁbroblasts (25). Ribose is the precursor of nucleic acids that is synthesized de novo through the nonoxidative pentose phosphate pathway using the thiamine-dependent enzyme transketolase (26). This hypothesis is consistent with the clinical ﬁndings of megaloblastic anemia in TRMA but awaits further studies to conﬁrm its validity. The relationship between a thiamine transport defect and diabetes mellitus is not well understood. While there are minimal data concerning thiamine and human pancreatic function, a series of experiments with thiamine-deﬁcient rats, performed by Sundaresen and colleagues, explored this issue (27–29). They documented that these rats developed hyperglycemia and had decreased insulin secretion basally and in response to several provocative agents. Proinsulin production in thiamine-deﬁcient rats was normal basally but decreased in response to a sulfonylurea, a known insulin secretagogue. Oxidative metabolism of glucose and pyruvate were decreased signiﬁcantly in isolated pancreatic islet cells compared to controls. Taken together, their work showed the production and secretion of insulin is abnormal when intracellular thiamine is inadequate in h-islet cells. The response of peripheral tissues does not appear to contribute to the etiology of diabetes mellitus in thiamine deﬁciency. First, the utilization of glucose by peripheral tissues does not decrease. In fact, studies of brain metabolism have revealed increased ﬂux through the glycolytic pathway in thiamine deﬁciency (30,31). In thiamine-deﬁcient rats that were hyperglycemic, the hypoglycemic response to insulin administration was normal, as was the response to an insulin secretagogue (32). It is unclear whether these studies with experimental thiamine deﬁciency are completely relevant for thiamine transport deﬁciency. These data seem at odds with some of the

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clinical observations concerning diabetes mellitus made with patients with TRMA. The constellation of medical problems associated with TRMA is reminiscent of those observed in several mitochondrial disorders. For example, TRMA overlaps substantially with Wolfram syndrome. Thus, it is interesting that Scharfe et al. in 2000 described a patient with TRMA and mitochondria dysfunction (33). She was a 14-year-old Turkish-German girl, the product of a consanguineous mating, who presented with short stature, hepatosplenomegaly, retinal degeneration, and abnormalities of the brain on MRI. While no mitochondrial abnormality was seen ultrastructurally, biochemical analysis showed a severe deﬁciency of pyruvate dehydrogenase complex and complex I of the respiratory chain. Lactate levels in serum and CSF were high. She was found to have inherited an SLC19A2 nonsense mutation in homozygosity. This study did not exclude the possibility that she had inherited a second genetic defect, perhaps attributable to consanguinity, which resulted in the mitochondrial disease. Song and Singleton in 2002 studied thiamine and thiamine pyrophosphate transport in mitochondria (34). Their studies documented that mitochondria isolated from TRMA lymphoblasts lacked the high-aﬃnity transport of thiamine observed in control mitochondria. This suggests that THT1 resides on both the cell and mitochondrial membranes. In contrast, the high-aﬃnity transport of thiamine pyrophosphate was preserved in TRMA mitochondria, consistent with previous studies establishing that this moiety uses a diﬀerent carrier (35). Since the primary thiamine species present intracellularly is thiamine pyrophosphate, it is unclear what eﬀect the loss of THT1 has on mitochondrial metabolism. In fact, the Km for the high-aﬃnity transport of thiamine in mitochondria is an order of magnitude higher than the intracellular concentration of thiamine. That said, it seems that THT1 must have some purpose in residing in the mitochondrial membrane, perhaps in transporting some other small organic molecule. The observation of striking ultrastructural abnormalities in TRMA erythroid cells (19) supports this suggestion. Iron-laden mitochondria are observed in a high proportion of the intermediate and late erythroblasts. The aﬀected mitochondria show degenerative changes, such as a loss of some or all of the cristae and the presence of abnormally electron-lucent areas in the mitochondrial matrix. Mitochondria also contain electron-dense intercristal deposits, which disappear after thiamine treatment. V. GENETICS, MAPPING, DISEASE GENE Mandel and colleagues described a 3-month-old girl who presented with severe megaloblastic anemia, diabetes, and sensorineural deafness, all of

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which improved with thiamine treatment (21). She was a child of parents who were ﬁrst cousins and also second cousins. Two maternal and two paternal aunts and uncles of the patients died early in life with severe anemia, sepsis, convulsions, and cardiorespiratory failure. In addition, Haworth and colleagues reported the occurrence of two cases of TRMA in one family (19). Review of these pedigrees implied that TRMA is inherited with an autosomal recessive mechanism. In 1997, Neufeld and coworkers performed linkage analysis with Alaskan and Italian kindreds inheriting TRMA (6). Since there was consanguinity in these families, they used a homozygosity mapping approach to link the trait to a region on chromosome 1q23.2-23.3 (maximum LOD score of 3.7 for DISI679). Haplotype analysis with additional consanguineous Israeli-Arab kindreds established a TRMA critical region spanning a 16 cM region. Further analyses by Raz et al. and Banikazemi et al. successively reﬁned the critical region to a more tractable size of 4 and 1.4 cM, respectively (7,8). Using high-throughout genomic sequences from the TRMA critical region, our group identiﬁed a putative novel gene with homology to SLC19A1, which encodes the reduced folate carrier 1(RFC1) (36–38). We cloned the entire coding region for this gene, named SLC19A2, from a human fetal brain cDNA library (11). This gene encodes 497-amino-acid protein predicted to have 12 transmembrane domains. We found two frameshift mutations in exon 2, a 1-bp insertion and a 2-bp deletion, which were predicted to obliterate the protein among four consanguineous Iranian families inheriting TRMA. Two other groups identiﬁed SLC19A2 as the TRMA disease gene in simultaneous publications. Nadine Cohen’s group used a positional cloning/ candidacy approach, constructing a PAC contig for the region (9). This resulted in further reﬁnement of the TRMA critical region to approximately 400 kb. They then studied genomic sequences from the same PAC that we had used and performed mutation analysis of all genes it contained, including SLC19A2. Ellis Neufeld’s group, using Cohen’s reﬁned region, identiﬁed SLC19A2 as a functionally relevant candidate (10). Both groups identiﬁed disease-causing mutations in SLC19A2. The SLC19A2 transcript is f3.8 kb in length and is expressed widely in human tissues, with most abundant steady-state levels in skeletal muscle and heart (9–12). The genomic organization consists of six exons that span approximately 22.5 kb (Fig. 1). As noted earlier, THT1 is highly homologous with RFC1 (39% amino acid identity and 57% similarity) with minimal homology at the N- and C-terminal regions as well as the loop between transmembrane helices VI and VII. When studied in vitro, it speciﬁcally transports thiamin in a Na+-independent manner (12), conﬁrming the prior studies of Rindi et al. and Stagg et al. (3–5). The transport

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Figure 1 Genomic organization of the SLC19A2 gene. The coding exons are shown as numbered ﬁlled boxes. The 5V and 3V untranslated regions are indicated with unﬁlled boxes. The positions of the ATG and stop codons are indicated. The scale in kilobases (kb) is indicated at the bottom.

of thiamin is H+ gradient dependent and is not inhibited by other organic cations. Fourteen SLC19A2 mutations have been reported to date (Fig. 2). (9– 11,18,30,39) Among them, ten mutations are nonsense defects that are predicted to truncate the protein prematurely. Those mutations were clustered between the sixth and seventh transmembrane domains. These trunca-

Figure 2 Distribution of SLC19A2 mutations causing TRMA. The predicted structure of the thiamine transporter, THT1, with 12 transmembrane domains is shown with the amino- and carboxy-terminal ends in the cytosolic compartment. The residues aﬀected by TRMA-causing mutations are indicated (ﬁlled circles) and the speciﬁc molecular lesions are noted. fs = frame shift.

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tion mutants are retained intracellularly (i.e., do not get transported to the cell membrane) (40). Three missense SLC19A2 mutations resulted in proteins that were not functional (41). Among these, two mutants were targeted to the cell membrane, while the third mutation appeared to result in an unstable transcript. The intracellular traﬃcking of the THT1 protein was characterized in 2002. The human SLC19A2 cDNA has two potential N-glycosylation sites and, indeed, the protein is N-glycosylated (41). Unlike some other transporters, however, this glycosylation does not appear to be important for its function as a transporter or the targeting of THT1 to the cell membrane. In contrast, THT1’s N-terminal sequence and transmembrane backbone do play an important role in the traﬃcking of the protein to the cell membrane, while the C-terminal domain is not required for this processing (40). Lastly, study of the dynamics of the traﬃcking of the vesicles containing THT1 showed that microtubles are involved in this intracellular transport but that microﬁlaments are not (40). VI.

AN ANIMAL MODEL OF TRMA

Our group and the Neufeld group independently cloned and characterized the orthologous mouse gene, Slc19a2 (13,42). This gene spans 16.3 kb and is organized into six exons, an organization that is conserved with the human SLC19A2. The Slc19a2 gene was localized to mouse chromosome 1, a region

Figure 3 Thiamine uptake by Slc19a2/ (diamonds), Slc19a2+/ (triangles) and Slc19a2+/+ (squares) erthyrocytes. (a) Ratio between the [3H]thiamine radioactivity in the cell (Ci) and in the medium (Cm). (b) Thiamine uptake of 25 AL of erythrocytes at diﬀerent concentration of thiamine in the medium. (Reprinted with permission from Oxford University Press.)

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Figure 4 Glucose tolerance, insulin secretion, and insulin tolerance testing. (a) GTT in Slc19a2/ (diamonds, n = 9) and Slc19a2+/+ (squares, n = 9) mice on thiamin-free diet at day 17. (b) IST in Slc19a2/ and Slc19a2+/+ mice on thiaminfree diet a day 14. Filled and open boxes indicate knockout (n = 3) and wild-type (n = 3) mice, respectively. (c) ITT in Slc19a2/ (diamonds, n = 2) and Slc19a2+/+ (squares, n = 4) mice on thiamin-free diet at day 16. Fasting glucose levels are 100% at 0 min. (Reprinted with permission from Oxford University Press.)

Figure 5 ABR thresholds in Slc19a2/ and Slc19a2+/+ mice. (a) Responses to click, 8-, 16-, and 32-kHz stimuli are shown. The open and ﬁlled boxes indicate regular and thiamine-free diets, respectively. (b) Waveforms for click stimulus on a 4-AV ﬁxed scale for comparison are shown. (Reprinted with permission from Oxford University Press.)

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syntenic to human chromosome 1q23 that contains SLC19A2 (42). Interestingly, Slc19a2 was also cloned using mRNA diﬀerential display from IW32 murine erythroleukemia cells expressing a temperature-sensitive p53 mutant allele,lwpVal-135, potentially providing new insights into the role of the thiamine transporter in cancer biology (43). The mouse protein product, Tht1, comprises 498 amino acid residues, is predicted to have 12 transmembrane domains, and shares a number of other conserved sequence motifs with the human orthologues. This includes one potential N-glycosylation site and several potential phosphorylation sites (13). Tht1 is a bona ﬁde thiamine transporter (42,43). It is expressed widely on the cell surface and intracellularly in mouse tissues. Speciﬁcally, it is localized to a subpopulation of cells in cochlea, small intestine, and pancreas (42), which have relevance for the TRMA phenotype. Recently, our group generated a mouse model of TRMA using targeted disruption of Slc19a2 through homologous recombination in embryonic stem cells (14). The high-aﬃnity component of thiamine transport was lacking in erythrocytes from the Slc19a2/ mice (Fig. 3). After 16 days of a thiaminefree dietary challenge, these mice developed diabetes mellitus, megaloblastosis, and sensorineural deafness. On glucose tolerance testing, the knockout mice showed severe fasting hyperglycemia and abnormal response to glucose loading with insuﬃcient insulin secretion (Fig. 4a and b). The response of serum glucose to exogenous insulin administration was normal (Fig. 4c), suggesting that peripheral tissues could utilize glucose properly. These data suggest that diabetes mellitus associated with TRMA results from inadequate insulin synthesis and/or secretion. No evidence of acute h-islet cell destruction was noted. Auditory evoked responses of the Slc19a2/ mice revealed normal hearing in mice maintained on standard mouse chow, but profound loss in those on a thiamine-free diet (Fig. 5). Of interest, repletion of thiamine in the diet resulted in signiﬁcant restoration of hearing in a majority of the animals tested. This ﬁnding provides support for the potential utility of thiamine therapy for preserving auditory function if initiated very early in life. ACKNOWLEDGMENTS This work was supported in part by awards from the NICHD (HD01294) and the March of Dimes (FY00-283) to B.D.G. REFERENCES 1.

LE Rogers, FS Porter, JBJ Sidbury. Thiamine-responsive megaloblastic anemia. J Pediatr 74:494–504, 1969.

TRMA: Clinical Aspects and Molecular Genetics 2.

3.

4.

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

561

MB Viana, RI Carvalho. Thiamine-responsive megaloblastic anemia, sensorineural deafness, and diabetes mellitus: a new syndrome? J Pediatr 93:235–238, 1978. G Rindi, D Casiorla, V Poggi, B De Vizia, C Patrini, U Laforenza. Thiamine transport by erythrocytes and ghosts in thiamine-responsive megaloblastic anaemia. J Inherit Metab Dis 15:231–242, 1992. 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. AR Stagg, JC Fleming, MA Baker, M Sakamoto, N Cohen, EJ Neufeld. Defective high-aﬃnity thiamine transporter leads to cell death in thiamineresponsive megaloblastic anemia syndrome ﬁbroblasts. J Clin Invest 103:723– 729, 1999. EJ Neufeld, H Mandel, T Raz, R Szargel, CN Yandava, A Stagg, S Faure, T Barrett, N Buist, N Cohen. Localization of the gene for thiamine-responsive megaloblastic anemia syndrome, on the long arm of chromosome 1 by homozygosity mapping. Am J Hum Genet 61:1335–1341, 1997. T Raz, T Barrett, R Szargel, H Mandel, EJ Neufeld, K Nosaka, MB Viana, N Cohen. Reﬁned mapping of the gene for thiamine-responsive megaloblastic anemia syndrome and evidence for genetic homogeneity. Hum Genet 103:455– 461, 1998. M Banikazemi, GA Diaz, P Vossough, M Jalali, RJ Desnick, BD Gelb. Localization of the thiamine-responsive megaloblastic anemia syndrome locus to a 1.4-cM region of 1q23. Mol Genet Metab 66:193–198, 1999. V Labay, T Raz, D Baron, H Mandel, H Williams, T Barrett, R Szargel, L McDonald, A Shalata, K Nosaka, S Gregory, N Cohen. Mutations in SLC19A2 cause thiamine-responsive megaloblastic anaemia associated with diabetes mellitus and deafness. Nat Genet 22:300–304, 1999. JC Fleming, E Tartaglini, MP Steinkamp, DF Schorderet, N Cohen, EJ Neufeld. The gene mutated in thiamine-responsive anaemia with diabetes and deafness (TRMA) encodes a functional thiamine transporter. Nat Genet 22: 305–308, 1999. GA Diaz, M Banikazemi, K Oishi, RJ Desnick, BD Gelb. Mutations in a new gene encoding a thiamine transporter cause thiamine-responsive megaloblastic anaemia syndrome. Nat Genet 22:309–312, 1999. B Dutta, W Huang, M Molero, R Kekuda, FH Leibach, LD Devoe, V Ganapathy, PD Prased. Cloning of the human thiamine transporter, a member of the folate transporter family. J Biol Chem 274:31925–31929, 1999. K Oishi, T Hirai, BD Gelb, GA Diaz. Slc19a2: cloning and characterization of the murine thiamin transporter cDNA and genomic sequence the orthologue of the human TRMA gene. Mol Genet Metab 73:149–159, 2001. K Oishi, S Hofmann, GA Diaz, T Brown, D Manwani, L Ng, R Young, H Vlassara, YA Ioannou, D Forrest, BD Gelb. Targeted disruption of Slc19a2, the gene encoding the high-aﬃnity thiamin transporter Thtr-1, causes diabetes

562

15.

16.

17.

18.

19. 20. 21. 22.

23.

24. 25.

26.

27.

28.

29.

Oishi et al. mellitus, sensorineural deafness and megaloblastosis in mice. Hum Mol Genet 11:2951–2960, 2002. FM Meire, MM Van Genderen, K Lemmens, MH Ens-Dokkum. Thiamineresponsive megaloblastic anemia syndrome (TRMA) with cone-rod dystrophy. Ophthalmic Genet 21:243–250, 2000. MR Abboud, D Alexander, SS Najjar. Diabetes mellitus, thiamine-dependent megaloblastic anemia, and sensorineural deafness associated with deﬁcient alpha-ketoglutarate dehydrogenase activity. J Pediatr 107:537–541, 1985. V Villa, A Rivellesse, F DiSalle, C Iovone, V Poggi, B Capaldo. Acute ischemic stroke in a young woman with the thiamine-responsive megaloblastic anemia syndrome. J Clin Endocrinol Metab 85:947–949, 2000. S Gritli, S Omar, E Tartaglini, S Guannouni, JC Fleming, MP Steinkamp, CI Berul, R Hafsia, SB Jilani, A Belhani, M Hamdi, EJ Neufeld. A novel mutation in the SLC19A2 gene in a Tunisian family with thiamine-responsive megaloblastic anaemia, diabetes and deafness syndrome. Br J Haematol 113:508–513, 2001. C Haworth, DI Evans, J Mitra, SN Wickramasinghe. Thiamine-responsive anaemia: a study of two further cases. Br J Haematol 50:549–561, 1982. A Bazarbachi, S Muakkit, M Ayas, A Taher, Z Salem, H Solh, JH Haider. Thiamine-responsive myelodysplasia. Br J Haematol 102:1098–1100, 1998. H Mandel, M Berant, A Hazani, Y Naveh. Thiamine-dependent beriberi in the ‘‘thiamine-responsive anemia syndrome.’’ N Engl J Med 311:836–838, 1984. G Valerio, A Franzese, V Poggi, A Tenore. Long-term follow-up of diabetes in two patients with thiamine-responsive megaloblastic anemia syndrome. Diabetes Care 21:38–41, 1998. V Poggi, G Rindi, C Patrini, B DeVizia, G Longo, G Andria. Studies on thiamine metabolism in thiamine-responsive megaloblastic anaemia. Eur J Pediatr 148:307–311, 1989. F Tenconi, M Bertoncelli, F Gatti, C Rosso. A novel vitamin B1 derivative: benzoyloxymethyl-thiamine (BT 851). Boll Chim Farm 122:27–44, 1983. EJ Neufeld, JC Fleming, E Tartaglini, MP Steinkamp. Thiamine-responsive megaloblastic anemia syndrome: a disorder of high-aﬃnity thiamine transport. Blood Cells Mol Dis 27:135–138, 2001. LG Boros, PW Lee, JL Brandes, M. Cascante, P Muscarella, WJ Schirmer, WS Melvin, EC Ellison. Nonoxidative pentose phosphate pathways and their direct role in ribose synthesis in tumors: Is cancer a disease of cellular glucose metabolism? Med Hypotheses 50:55–59, 1998. P Rathanaswami, A Pourany, R Sundaresan. Eﬀects of thiamine deﬁciency on the secretion of insulin and the metabolism of glucose in isolated rat pancreatic islets. Biochem Int 25:577–583, 1991. P Rathanaswami, R Sundaresan. Modulation of A and B cell functions by tolbutamide and arginine in the pancreas of thiamine-deﬁcient rats. Biochem Int 19:793–802, 1989. P Rathanaswami, R Sundaresan. Eﬀects of thiamine deﬁciency on the biosynthesis of insulin in rats. Biochem Int 24:1057–1062, 1991.

TRMA: Clinical Aspects and Molecular Genetics

563

30. T Murata, N Omata, Y Fujibayashi, A Waki, N Sadato, M Yoshimoto, M Omori, K Isaki, Y Yonekura. Dynamic changes in glucose metabolism induced by thiamine deﬁciency and its replenishment as revealed by a positron autoradiography technique using rat living brain slices. J Neurol Sci 164:29–36, 1999. 31. AM Hakim, HM Pappius. Sequence of metabolic clinical and histological events in experimental thiamine deﬁciency. Ann Neurol 13:365–375, 1983. 32. P Rathanaswami, R Sundaresen. Eﬀects of insulin secretagogues on the secretion of insulin during thiamine deﬁciency. Biochem Int 17:523–528, 1988. 33. C Scharfe, M Hauschild, T Klopstock, AJ Janssen, PH Heidemann, T Jaksch, M Jaksch. A novel mutation in the thiamine-responsive megaloblastic anaemia gene SLC19A2 in a patient with deﬁciency of respiratory chain complex I. J Med Genet 37:669–673, 2000. 34. Q Song, CK Singleton. Mitochondria from cultured cells derived from normal and thiamine-responsive megaloblastic anemia individuals eﬃciently import thiamine diphosphate. BMC Biochem 3:8, 2002. 35. R Zhao, F Gao, Y Wang, GA Diaz, BD Gelb, ID Goldman. Impact of the reduced folate carrier on the accumulation of active thiamin metabolites in murine leukemia cells. J Biol Chem 276:1114–1118, 2001. 36. PD Prasad, S Ramamoorthy, GH Leibach, V Ganapathy. Molecular cloning of the human placental folate transporter. Biochem Biophys Res Commun 206:681–687, 1995. 37. FM Williams, WF Flintoﬀ. Isolation of a human cDNA that complements a mutant hamster cell defective in methotrexate uptake. J Biol Chem 270:2987– 2992, 1995. 38. SC Wong, SA Proefke, A Bhushan, LH Matherly. Isolation of human cDNAs that restore methotrexate sensitivity and reduced folate carrier activity in methotrexate transport-defective Chinese hamster ovary cells. J Biol Chem 270:17468–17475, 1995. 39. T Raz, V Labay, D baron, R Szargel, Y Anbinder, T Barrett, W Rabl, MB Viana, H Mandel, A Baruchel, JM Cayuela, N Cohen. The spectrum of mutations, including four novel ones, in the thiamine-responsive megaloblastic anemia gene SLC19A2 of eight families. Hum Mutat 16:37–42, 2000. 40. VS Subramanian, JS Marchant, I Parker, HM Said. Cell biology of the human thiamine transporter-1 (hTHTR1): intracellular traﬃcking and membrane targeting mechanisms. J Biol Chem, e-pub, 2002. 41. K Balamurugan, HM Said. Functional role of speciﬁc amino acid residues in human thiamine transporter SLC19A2: mutational analysis. Am J Physiol Gastrointest Liver Physiol 283:G37–43, 2001. 42. JC Fleming, MP Steinkamp, R Kawatsuji, E Tartaglini, JL Pinkus, MD Fleming, EJ Neufeld. Characterization of a murine high-aﬃnity thiamine transporter Slc19a2. Mol Genet Metab 74:273–280, 2001. 43. PK Lo, JY Chen, PP Tang, J Lin, CH Lin, LT Su, CH Wu, TL Chen, Y Yang, FF Wang. Identiﬁcation of a mouse thiamine transporter gene as a direct transcriptional target for p53. J Biol Chem 276:37186–37193, 2001.

32 Accomplishments and Future Directions Frank Jordan Rutgers University, Newark, New Jersey, U.S.A. Mulchand S. Patel School of Medicine and Biomedical Sciences, State University of New York at Buﬀalo, Buﬀalo, New York, U.S.A.

The ﬁrst half of the volume is devoted to studies of catalytic mechanisms of enzymes, simpler than the a-keto acid dehydrogenase complexes discussed later. One of the important merits of these contributions is the interweaving of structural and chemical/biochemical information. After a historical introduction, current knowledge on the biosynthesis of thiamine itself by bacteria and of thiamine diphosphate by both a bacterial and mammalian source is described. A striking feature of the latter is that mammalian cells convert thiamine directly to thiamine diphosphate, while in bacteria the ﬁrst thiamine phosphate is the monophosphate. The structures for the enzymes pyruvate decarboxylase, benzoylformate decarboxylase, acetohydroxyacid synthase, transketolase, along with mechanistic ﬁndings on each were next presented. The mechanisms of these enzymes are being evaluated in greater and greater detail, using a number of ingenious methods to establish the existence of enzyme-bound intermediates, all of which are covalent thiamine diphosphate-bound conjugates. Notable among the ﬁndings are: crystallographic detection of the key 565

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enamine intermediate on transketolase; proton nuclear magnetic resonance detection of intermediates produced in a rapid chemical quench system on several enzymes, enabling the authors to determine the rate-limiting step on those enzymes; visible spectroscopic determination of the kinetic fate of two consecutive intermediates on benzoylformate decarboxylase. The authors also made great strides assigning roles to active-center residues of all of these enzymes. Such assignments are quite diﬃcult, and from the reports two general conclusions have emerged. Even for two enzymes carrying out very similar reactions, such as pyruvate decarboxylase and benzoylformate decarboxylase, the acid–base groups at the active center are not the same in the corresponding positions or for the corresponding function. Also, there is more evidence that several active-center residues may participate in catalysis in more than one elementary reaction step. On the basis of a variety of experimental ﬁndings—including the conservation of the ‘‘V’’ coenzyme conformation and of the three highly conserved hydrogen bonds to the 4V-aminopyrimidine ring of the coenzyme; the recent direct detection at 305 nm of the absorbance pertinent to the imino tautomer of the coenzyme both on yeast pyruvate decarboxylase and on the E. coli pyruvate dehydrogenase complex E1 component—it is now safe to insert this imino tautomer on all of the pathways using thiamine diphosphate. This ﬁnding makes this cofactor unique so far in being capable of providing both electrophilic and acid–base catalytic functions for the reactions. In addition to the structure–function mechanistic studies, there were several important summaries on the use of thiamine diphosphate–dependent enzymes in the chiral synthesis of a number of pharmacologically important intermediates. Classical among these is the formation of a-hydroxyketones with great enantiomeric excess. This is a growing ﬁeld and, as some of the contributors so elegantly demonstrated already, these enzymes can carry out condensations with a variety of other functional groups as well. With all of these potential synthetic applications, these enzymes are bound to become important synthetic tools in the chemical/pharmaceutical industry. In the second half of this volume, the family of the multienzyme a-keto acid dehydrogenase complexes involved in the thiamine diphosphate– dependent oxidation of pyruvate, branched-chain a-keto acids and a-ketoglutarate in the mitochondria that play a central role in oxidative metabolism in higher organisms is discussed. These complexes constitute some of the largest and most complex enzyme systems, and two of them are controlled by dedicated and intricately regulated phosphorylation–dephosphorylation systems. In the recent past, there have been several important developments in various aspects of the structure–function relationship of component proteins, regulation, the characterization of the genes, and their involvement in inherited metabolic disorders. Protein engineering approaches have sup-

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ported biophysical studies such as electron cryomicroscopy, X-ray crystallography and NMR spectroscopy that have produced detailed three-dimensional structures providing important insights at the atomic level relevant to the details of catalytic mechanisms and the molecular basis of a spectrum of genetic disorders. In Section IV entitled ‘‘Structure–Function of Thiamine-Diphosphate Multienzyme Complexes,’’ a series of chapters discussed the aspects of the molecular architecture of subcomplexes (component–component interactions), assembly of the component structures, active-site coupling and substrate channeling involving ﬂexible lipoyl domains, and the catalytic mechanisms by the components of these multienzyme complexes. Studies of structural dynamics have revealed that ‘‘breathing’’ of the pyruvate dehydrogenase complex-E2 core is an intrinsic property and may contribute to its function. Evidence is also presented for the central role of the lipoyl domains not only for active-site coupling but also in providing structural interactions for the regulatory components (speciﬁc kinases and phosphatases) in higher eukaryotic pyruvate dehydrogenase complexes. The crystal structure of the pyruvate dehydrogenase complex-E1 (the a2 homodimeric form) from E. coli has provided structure–function relationships in this homodimeric enzyme compared with a heterotetrameric branched chain a-ketoacid dehydrogenase complex-E1 requiring thiamine diphosphate as a cofactor. Additionally, identiﬁcation of several residues in the active site of the pyruvate dehydrogenase complex-E1 provides in-depth analysis of the mechanism of catalysis and involvement of a speciﬁc histidyl residue in catalysis. Similarly, the crystal structure of a a2h2 heterotetrameric E1 component of human branched-chain a-ketoacid dehydrogenase complex has revealed the subunit–subunit interactions and the interactions of thiamine diphosphate with speciﬁc amino acid residues in the active site. For the lipoyl moiety to participate in catalysis its activation and covalent attachment to a speciﬁc lysyl residue(s) of E2s is a prerequisite. The requirement of one or more enzymes to fulﬁll this function in bacteria and higher eukaryotic organisms has shed light on evolutionary diversion. The three-dimensional structure of mammalian pyruvate dehydrogenase kinase 2 and branchedchain a-ketoacid dehydrogenase kinase provided important information on the functional domains and their roles in the catalysis. Interestingly, pyruvate dehydrogenase kinase 2 is remarkably similar to the structure of bacterial histidine kinases; however, its catalytic mechanism is similar to that of serine kinases. Further studies on the interactions of the components of these complexes promise to provide insight to both intersubunit information transfer and assembly of the entire complex. The section on Biomedical Aspects of Thiamine Diphosphate– Dependent Enzymes covers a spectrum of diseases, either genetic or acquired,

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associated with thiamine phosphate–requiring enzymes. Genetic defects in the pyruvate dehydrogenase complex and branched-chain a-ketoacid dehydrogenase complex resulting in neurological dysfunction and beneﬁcial eﬀects of thiamine therapy in a limited number of aﬀected patients have been discussed in detail. Furthermore, the biochemical basis for the impaired function for several speciﬁc mutations have been presented in light of the three-dimensional structure of E1 components. Neuropathologic ﬁndings in the brain from patients with the Wernicke– Korsakoﬀ syndrome (due to thiamine deﬁciency) are summarized, and the possible role of increased endothelial isoform of nitric oxide synthase in a-ketoglutarate dehydrogenase complex is implicated, suggesting a common pathophysiological mechanism for neurodegenerative diseases. Furthermore, exciting new developments using an animal model for thiamine-responsive megaloblastic anemia syndrome provided insights into the pathogenesis of diabetes mellitus in patients with this syndrome. It is clear from the contributions in this volume that the ﬁeld of thiamine diphosphate enzymes remains a very vibrant and productive one, with new members being discovered all the time. At the same time, the recently solved high-resolution structures of the physiologically relevant members of the family have provided an unprecedented opportunity to elucidate the structural/functional origins of diseases produced by mutations in the genes from which these enzymes are derived.

Index

Acetohydroxyacid synthase, 251–274 acetolactate, competition between, acetohydroxybutyrate formation, 238 active site of AHAS II, in process of condensation of 2-ketobutyrate with hydroxyethyl thiamine diphosphate, 234, 235 active sites, 263–265 bacterial, 233–250 beta-domain of yeast AHAS containing FAD, 261, 263 branched-chain amino acid biosynthesis, 251–252 catalytic subunit monomer of yeast AHAS, structure of, 259, 260 cofactors, 252–253 in yeast AHAS, POX, locations, conformations of, 266 covalent intermediates, 68, 69 distribution, 253 enzymatic characterization, 254–259 Escherichia coli, 254–255 regulation of, 256–259

[Acetohydroxyacid synthase] in vitro yeast acetohydroxyacid synthase reconstitution, 255–256 FAD, 264–265 herbicidal inhibitors, 251, 252 herbicide-binding site, 265–269 herbicide-resistance mutations, 265–268 natural role of, 269 structure of, 268–269 yeast AHAS, 267, 268 location of cofactors in, 263–265 minimal mechanism for, 239, 240 potassium phosphate concentration, eﬀect on yeast AHAS, 255, 256 properties of, 237 reactions catalyzed by, 233, 234, 251, 252 reconstituted yeast AHAS, regulation of, 256, 257 residues contacting thiamine diphosphate, in yeast AHAS, 264 569

570 [Acetohydroxyacid synthase] R-phenyl acetyl carbinol, reaction of pyruvate, benzaldehyde in foration of, 241, 242 S-acetolactate, AHAS R, stereochemistry of formation of, 241, 242 sources of, 253 structure, 259–265 subunits, 253–254 thiamine diphosphate, 67–68, 264 thiamine diphosphate-dependent enzymes: C alpha atoms, overlay of, 263 comparison with, 261–263 three-dimensional structure of, 259–261 yeast acetohydroxyacid synthase: crystallization of, 259 three-dimensional structure of, 259–261 Acetohydroxybutyrate, acetolactate formation, competition between, bacterial acetohydroxyacid synthases, 238 Acetoin-ribose-5-phosphate transaldolase, 94 Acetolactate, acetohydroxybutyrate formation, competition between, bacterial acetohydroxyacid synthases, 238 Acetolactate synthase, 94 Acetyl-thiamine diphosphate, in aqueous solutions, structures of, 9, 10 Acyl-GMP formation, lipoic acid in protein lipoylation1, catalyzed by, 348, 350 Acyloin condensations, carbon-carbon bonding reactions, thiamine-dependent enzymes as catalysts, ‘‘Orphan’’ thiamine diphosphate decarboxylases, 78

Index Acyl-thiamine diphosphate intermediates, 3, 4–11 enzymatic intermediate, identiﬁcation of as, 9–11 ﬂuoropyruvate, as alternative substrate leading to, 7–8 in reactions of alpha-ketoacid dehydrogenase complexes, 5–7 synthesis of, 8–9 thiamine diphosphate catalysis, role of, 3, 5 Adrenal gland, thiamine triphosphatase, 53 Alpha-carbanion/enamine reaction intermediate in transketolase site, kinetic crystallography, 159–172 beta hydroxylpyruvic acid binding to transketolase, circular dichroism measurement, kinetics of, 162–163 circular dichroism measurements, 161 conformational changes, intermediate formation and, 167–168 crystallographic model building, reﬁnement, 162 crystallography, 161 data collection, 161 dihydroxy ethyl-thiamine diphosphate, interactions with active-site residues, 165–167 dihydroxy ethyl-thiamine diphosphate intermediate, 164–165 near-UV CD spectra reﬂecting formation of, 167 D,L-2-(1,2-dihydroxy ethyl)thiamine diphosphate in active site of transketolase, surroundings of reaction intermediate, 166

Index [Alpha-carbanion/enamine reaction intermediate in transketolase site, kinetic crystallography] election density maps, 163 formation of, using beta hydroxypyruvic acid as donor substrate, 160 protein expression, puriﬁcation, 161 pyruvate, ferredoxin oxidoreductase, 168, 169 transketolase, ferredoxin oxidoreductase, 168, 169 transketolase-beta hydroxylpyruvic acid adduct, kinetics of formation of, 161 Alpha-ketoacid dehydrogenase complex: acyl-thiamine diphosphates, in reactions of, 5–7 mammalian branched-chain, 449–450 crystal structure, 450–462 lipoyl-bearing domain, solution structure, dynamics, 462–465 maple syrup urine disease, 450–458 thiamine diphosphate: catalysis, 6 reaction mechanisms, 6 Alpha-(mandelyl)thiamine intermediate, in thiaminecatalyzed decarboxylation of benzolformate, 299–300 Alpha-oxidation pathway, 2hydroxyphytanoyl-COa lyase, thiamine-dependent enzyme, 530–534 Amygdala, thiamine triphosphatase, 53 Anemia, megaloblastic, thiamineresponsive, 549–564 animal model, 557–560 biochemical aspects, 552–554 clinical manifestations, 550–551 genetics, 554–557 treatment, 551–552

571 Antibiotic bacimethrin, conversion to methoxythiamine pyrophosphate, by thiamine pyrophosphate biosynthetic enzymes, 24 Aorta, thiamine triphosphatase, 53 Apex of heart, thiamine triphosphatase, 53 Apnea, with pyruvate dehydrogenase complex deﬁciency, 473 Apotransketolase, holotransketolase, induced absorption band of, 146, 147 Appendix, thiamine triphosphatase, 53 Asymmetric cross-benzoin condensation, with benzaldehyde lyase, 125–126 Ataxia, with pyruvate dehydrogenase complex deﬁciency, 473 Atrium, thiamine triphosphatase, 53 Bacillus stearothermophilus, pyruvate dehydrogenase multienzyme complex, 332, 333 Bacimethrin toxicity, thiamine biosynthetic enzymes, 23–24 Bacterial acetohydroxyacid synthases, 233–250 acetohydroxyacid synthase II, properties of, 237 acetolactate, acetohydroxybutyrate formation, competition between, 238 active site of acetohydroxyacid synthase II, in process of condensation of 2-ketobutyrate with hydroxyethyl thiamine diphosphate, 234, 235 branched-chain amino acids, feedback inhibition by, 242–247 C-terminal domain of regulatory subunit, 245–247 valine site structure, 243–245

572 [Bacterial acetohydroxyacid synthases] catalysis, 234–242 methionine 250, 238–239 minimal mechanism for, 239, 240 new condensation reaction catalyzed by, 241–242 polar side chains, 239–241 reaction catalyzed by, 233, 234 R-phenyl acetyl carbinol: acetohydroxyacid synthase II, stereochemistry of formation of, 241, 242 reaction of pyruvate, benzaldehyde in formation of, 241, 242 S-acetolactate, acetohydroxyacid synthase II, stereochemistry of formation of, 241, 242 speciﬁcity, 234–242 Trp464, properties of, 237 Val375, properties of, 237 Val461a, properties of, 237 Bacterial thiamine biosynthetic pathway, 15, 16 Benzaldehyde lyase: active-site residues, benzoylformate decarboxylase, 280, 282 assay of, benzoylformate decarboxylase, 279 asymmetric cross-benzoin condensation with, 125–126 benzoylformate decarboxylase, active-site residues, 280, 282 model of active site of, 280, 282 pyruvate decarboxylase, active-site residues, 280, 282 reactions catalyzed by, 122–126, 275, 276 Zymomonas mobilis, active-site residues, 280, 282 Benzaldehyde lyase variants, benzoylformate decarboxylase, construction, expression, puriﬁcation of, 278–279

Index Benzaldehyde lyase-catalyzed carbon-carbon bond formation: enantioselective synthesis of hydroxy ketones, 113–130 hydroxy ketones, enantioselective synthesis, 113–130 Benzaldehyde lyase-mediated benzoin condensation, 124 Benzaldehyde lyase-mediated carboligation, (R)-2-HPP derivatives, 125 Benzoin aldolase, 94 Benzolformate, thiamine-catalyzed decarboxylation, alpha(mandelyl)thiamine intermediate, 299–300 Benzoylformate decarboxylase, 94, 131–142, 275–290, 291–308 alpha-(mandelyl)thiamine intermediate, in thiaminecatalyzed decarboxylation of benzolformate, 299–300 benzaldehyde lyase, assay of, 279 benzaldehyde lyase variants, construction, expression, puriﬁcation of, 278–279 benzoylformate decarboxylase variants, construction, expression, puriﬁcation of, 277–278 buﬀer changes, 302–306 buﬀer concentration, absorbance of solutions of mandelythiamin, change with, 302, 303 crystal form I, crystal contact between tetramers in, 134 crystal form V, binding of Rmandelate to active site in, 136, 137 crystal forms, comparison of, 133 decarboxylase activity, assay of, 279 decarboxylation of alpha(mandelyl)thiamine, 300–301

Index [Benzoylformate decarboxylase] enzymatic roles of residues, 135–139 kinetic studies, 137–139 structural studies, 136–137 fragmentation products, 301 his-tagged PDC variants, construction, expression, puriﬁcation of, 277 hydroxybenxyl thiamine: benzaldehyde production, 293–295 C2alpha proton exchange, 293 fragmentation of, 296 hydroxybenzyl thiamine, 292–293 hydroxybenzyl thiamine yield, as function of buﬀer concentration, 304 intermediates in, 291–292 N-alkylation of hydroxybenxyl thiamin, charge localization, 295 phosphate bugger concentration, rate of fragmentation, 297, 298 reactions catalyzed by, 275, 276 roles of active-site residues, 135 saturation, eﬀect of buﬀer subject to, 296–299 Ser26, 139–140 structure, 132–134 active site in crystal form I, form V, R-mandelate bound, 132, 133 wild-type: benzoin condensation, to benzoins, mediation, 116 carboligation, mediated, toward (S)-2-HPP derivatives, 114 catalyzed by carbon-carbon bond formations, control of enantioselectivity in, 116 mutants, formation of (S)-2HPPalpha, 122 Benzoylformate decarboxylase H281A, asymmetric cross-benzoin condensation with, 119–120 Benzoylformate decarboxylase muteins, 114–122

573 Benzoylformate decarboxylase variants: construction, expression, puriﬁcation of, 277–278 with high carboligation activity, in presence of water-miscible organic solvents, 120–121 as solution of ‘‘ortho-problem,’’ 121–122 Benzoylformate decarboxylasecatalyzed carbon-carbon bond formation: enantioselective synthesis of hydroxy ketones, 113–130 hydroxy ketones, enantioselective synthesis, 113–130 Benzoylformate decarboxylasemediated carboligation, wild type, toward (S)-2-HPP derivatives, 114 Bladder, thiamine triphosphatase, 53 Bone marrow, thiamine triphosphatase, 53 Brain: malformation of, with pyruvate dehydrogenase complex deﬁciency, 473 thiamine triphosphatase, 53 Branched-chain alpha-ketoacid dehydrogenase: mammalian complex, 449–450 crystal structure, 450–462 lipoyl-bearing domain, solution structure, dynamics, 462–465 maple syrup urine disease, 450–458 maple syrup urine disease, 510–512, 512–513, 517–518 thiamine responsiveness, 510 regulation of, 510–512 Branched-chain alpha-ketoacid dehydrogenase E1 component, 34, 36 Branched-chain amino acid biosynthesis, acetohydroxyacid synthase, 251–252

574 Burkitt’s lymphoma, thiamine triphosphatase, 53 Carbon kinetic isotope eﬀects, yeast pyruvate decarboxylase, 217–232 carbon isotope eﬀects, 224 isotope eﬀects, on kcat for yeast pyruvate decarboxylase variants, 224–226 kinetic isotope eﬀects, transition-state theory, 218–219 low pyruvate concentrations, for pyruvate decarboxylase variants, isotope eﬀects at, 226–228 solvent isotope eﬀects, 223–224 steady-state kinetics, variants, 220–222 wild-type yeast pyruvate decarboxylase reaction, rate-limiting steps, 220 Carbon-carbon bond cleavage, kinetic racemic resolution via, 123–125 Carbon-carbon bond formation: catalyzed by wild-type benzoylformate decarboxylase, control of enantioselectivity in, 116 enantioselective synthesis of hydroxy ketones, benzaldehyde lyase, benzoylformate decarboxylase-catalyzed, 113–130 Carbon-carbon bonding reactions: ‘‘Orphan’’ thiamine diphosphate decarboxylases, 77–93 acyloin condensations, 78 C-1 metabolism, 82–84 imines as acceptors, 84–86 ketones as acceptors, 84–86 range of reactions catalyzed by, 79–82

Index [Carbon-carbon bonding reactions] thiamine-dependent enzymes as catalysts, 77–93 acyloin condensations, 78 C-1 metabolism, 82–84 imines as acceptors, 84–86 ketones as acceptors, 84–86 range of reactions catalyzed by ‘‘orphan’’ thiamine diphosphate decarboxylases, 79–82 Cardiovascular tissue, thiamine triphosphatase, 53 Catalysis: pyruvate dehydrogenase multienzyme complex, 339–340 thiamine in enzymes, model of, 66 Caudate nucleus, thiamine triphosphatase, 53 Central nervous system degeneration, with pyruvate dehydrogenase complex deﬁciency, 473 Cerebellum, thiamine triphosphatase, 53 Cerebral cortex, thiamine triphosphatase, 53 Clontech, human multiple-tissueexpression array, thiamine triphosphatase, mRNA distribution, human tissue, 53 Colon, thiamine triphosphatase, 53 Colorectal adenocarcinoma, thiamine triphosphatase, 53 Congenital brain malformation, with pyruvate dehydrogenase complex deﬁciency, 473 Corpus callosum, thiamine triphosphatase, 53 Covalent intermediates: in enzymic thiamine catalysis, 61–72 superfamily of thiamine diphosphate-dependent enzymes, 68, 69 thiamine in enzymes, 68, 69

Index Crystal soaking, thiamine pyrophosphokinase, 30 Crystallization, thiamine pyrophosphokinase, 30 1-deoxy-D-xylulose 5-phosphate synthase, 94 Developmental delay, with pyruvate dehydrogenase complex deﬁciency, 473 Digestive system, thiamine triphosphatase, 53 Dihydrolipoamide: acetyltransferase, 485, 487, 500, 502 (See also E2 within text): E1-binding domain of, 485 inner domains of, 485 lipoyl domains of, 485, 498, 500 lipoyl moieties of, 498, 502 dehydrogenase, 485, 487 (See also E3 within text): binding protein, 485 Duodenum, thiamine triphosphatase, 53 Dysmorphic features, with pyruvate dehydrogenase complex deﬁciency, 473 Enamine-thiamine diphosphate, thiamine diphosphate catalysis, role of, 3, 5 Enantioselective synthesis, hydroxy ketones: benzaldehyde lyase, benzoylformate decarboxylase-catalyzed carbon-carbon bond formation, 113–130 benzaldehyde lyase-carbon-carbon bond formation, 113–130 Escherichia coli: acetohydroxyacid synthase, enzymatic characterization, 254–255 lipoic acid in protein lipoylation1, metabolism of lipoic acid in, 357

575 [Escherichia coli] pyruvate dehydrogenase multienzyme complex, information transfer, 407–432 407-residue, 418–422 active-site environment, 409–411 cofactor binding, 409–411 on Escherichia coli PDHc-E1, mechanistic information, 413–429 H179 residue, 414–418 histidine residues, in active center, 413–414 negative circular dichroism band, near 320 nm, 427–429 PDHc-E1 reaction, 426–427 phosphonolactyl thiamine diphosphate, iminopyrimidine tautomeric form of, 422–426 related enzymes, structural comparison with, 411–413 structural information, on Escherichia coli PDHc-E1, 409–413 subunit assembly, 409 Y177 residue, 414–418 replacement: pyruvate dehydrogenase complex of: aceE-IpdA region, restructuring, operon, 393–395 changing coenzyme speciﬁcities, PDH, ODH complexes, 389–393 genetically engineered variants, 387–406 lipoyl domains, per acetyl transferase chain, 395–399 pyruvate oxidase, replacing PDH complex by, physiological consequences of, 399–401 pyruvate oxidase, pyruvate dehydrogenase complex of, 387–406 Esophagus, thiamine triphosphatase, 53

576 Ethanol, eﬀect on thiamine pyrophosphokinase, 32, 33 Exon-intron organization, human lipoyltransferase gene, lipoic acid in protein lipoylation1, 355 Ferredoxin oxidoreductase: alpha-carbanion/enamine reaction intermediate, transketolase site, kinetic crystallography, 168, 169 thiazolium ring structures, in reaction intermediates, 168, 169 Fetal tissue, thiamine triphosphatase, 53 Fluoropyruvate: acyl-thiamine diphosphates, as alternative substrate leading to, 7–8 pyruvate dihydrogenase complex, substrate for E1 component of, 8 Formaldehyde transketolase, 94 Free-radical reactions, role of thiamine diphosphate, 11–12 Frontal lobe, brain, thiamine triphosphatase, 53 Fructose-6-phosphate phosphoketolase, 94 Growth retardation, with pyruvate dehydrogenase complex deﬁciency, 473 5-guanidino-2-oxopentanoate decarboxylase, 94 Heart, thiamine triphosphatase, 53 Herbicidal inhibitors, acetohydroxyacid synthase, 251, 252 Herbicide-binding site: acetohydroxyacid synthase, 265–269 herbicide-resistance mutations, 265–268 natural role of, 269 structure of, 268–269 yeast acetohydroxyacid synthase, 267, 268

Index Herbicide-resistance mutations, acetohydroxyacid synthase, 265–268 Hippocampus, thiamine triphosphatase, 53 Holotransketolase, induced absorption band of, 143–158 aminopyrimidine ring of thiamine diphosphate in, hydrophobic pocket of, 145 apotransketolase, beta hydroxypyruvic acid, absorption spectra, in presence, absence, 146, 147 beta hydroxypyruvic acid, absorption spectra, in presence, absence, 146, 147 equilibrium formation, as function of thiamine diphosphate concentration, 146, 148 intensity of thiamine diphosphate absorption band, acidity of medium and, 152, 153 single turnover of beta hydroxypyruvic acid conversion by, 148, 149 thiamine diphosphate, beta hydroxypyruvic acid, absorption spectra, in presence, absence, 146, 147 Hydroxy ketones, enantioselective synthesis of: benzaldehyde lyase- and benzoylformate decarboxylase-catalyzed carbon-carbon bond formation, 113–130 benzaldehyde lyase-catalyzed carbon-carbon bond formation, 113–130 benzoylformate decarboxylase carbon-carbon bond formation, 113–130 benzoylformate decarboxylase-catalyzed carbon-carbon bond formation, 113–130 meta-substituted benzaldehydes, 118, 119 para-substituted benzaldehydes, 118

Index 2-hydroxy-3-oxoadipate synthase, 94 Hydroxybenxyl thiamine: benzaldehyde production, 293–295 C2alpha proton exchange, 293 fragmentation of, 296 N-alkylation of, 295 Hydroxybenzyl thiamine, 292–293 Hydroxybenzyl thiamine yield, as function of buﬀer concentration, benzoylformate decarboxylase, 304 Hydroxyethyl thiamine diphosphate, 204–205 2-hydroxyphytanoyl-COa lyase, thiamine-dependent enzyme, 532–536 Hypotonia, with pyruvate dehydrogenase complex deﬁciency, 473 Hypoventilation, with pyruvate dehydrogenase complex deﬁciency, 473 Ileum, thiamine triphosphatase, 53 Ilocecum, thiamine triphosphatase, 53 Immune system, thiamine triphosphatase, 53 Indolpyruvate decarboxylase, 94 Induced absorption band, holotransketolase, 143–158 Interventricular septum, thiamine triphosphatase, 53 Jejunum, thiamine triphosphatase, 53 Ketomalonate, least squares alignment of structures, yeast pyruvate decarboxylase, 178, 179 Kidney, thiamine triphosphatase, 53 Kinetic crystallography, alpha-carbanion/enamine reaction intermediate in transketolase site, 159–172 beta hydroxylpyruvic acid binding to transketolase, circular dichroism measumement, kinetics of, 162–163

577 [Kinetic crystallography, alpha-carbanion/enamine reaction intermediate in transketolase site] circular dichroism measurements, 161 conformational changes, intermediate formation and, 167–168 crystallographic model building, reﬁnement, 162 crystallography, 161 data collection, 161 dihydroxy ethyl-thiamine diphosphate, interactions with active-site residues, 165–167 dihydroxy ethyl-thiamine diphosphate intermediate, 164–165 near-UV CD spectra reﬂecting formation of, 167 election density maps, 163 protein expression, puriﬁcation, 161 transketolase-beta hydroxylpyruvic acid adduct, kinetics of formation of, 161 Lactobacillus plantarum, ligandinduced conformational changes, 95, 96 pyruvate oxidase, 99–102 Lactyl-thiamine diphosphate, 203–204 Leucocytes, peripheral, thiamine triphosphatase, 53 Leukemia, thiamine triphosphatase, 53 Lipoic acid, in protein lipoylation, 343–362 acyl-CoA synthetase, reaction catalyzed by LAE/medium-chain, 350, 351 acyl-GMP formation, catalyzed by, 348, 350 cDNA, deduced amino acid sequence, nucleotide sequence of, 348, 349 Escherichia coli, metabolism of lipoic acid in, 357

578 [Lipoic acid, in protein lipoylation] exon-intron organization, human lipoyltransferase gene, 355 lipoate acceptor proteins, lipoyltransferase mRNA, mRNAs of, 356 lipoate-activating enzyme, puriﬁcation, characterization of, 344–348 lipoic acid: activation of, 344–351 amino acid sequences surrounding attachment site, 352 lipoyltransferase: cDNA structure, 353–356 properties of, 351–353 lipoyltransferases, Escherichia coli lipoate-protein ligase A, 353, 354 mitochondria, metabolism of lipoic acid in, 357 mitochondrial medium-chain acylCoA synthetase, lipoate-activating enzyme, 348–351 proteins, transfer of lipoic acid to, 351–356 reaction products, 346, 347 structure, 343, 344 Lipoly-lysine residue, lipoyl domain, pyruvate dehydrogenase multienzyme complex, 334, 335 Lipoyl domain, pyruvate dehydrogenase multienzyme complex: interaction with E1, 334–337 reaction mechanism, 334–337 Lipoyl domain-mediated activated function, control, pyruvate dehydrogenase complex, PD kinases, phosphatase 1, 363–386 composition, 363–364 E2-domain structures, domain roles, 366–368 E2-facilitated kinase function, 370– 377 kinase activity, E2-mediated, modiﬁed regulation of, 373–377

Index [Lipoyl domain-mediated activated function, control, pyruvate dehydrogenase complex, PD kinases, phosphatase 1] organization of, 368–370 PDK2, binding, activation of, 370– 372 PDK3, binding, activation of, 372– 373 PDK4, binding speciﬁcity, regulation, 377 PDP1, E2-mediated CA2+ activation of, 377–379 regulatory enzymes, 364–365 Lipoyltransferase: Escherichia coli lipoate-protein ligase A, 353, 354 genomic DNA structure, 353–356 properties of, 351–353 Liver, fetal, thiamine triphosphatase, 53 Lung, carcinoma of, thiamine triphosphatase, 53 Maple syrup urine disease, 450–458 thiamine responsiveness, 509–524 branched-chain alpha-ketoacid dehydrogenase, 510 regulation of, 510–512 by complex-speciﬁc kinase, 512–513 role of thiamine pyrophosphate in, 517–518 history of disease, 509–510 model systems, 518–519 mutations, 514–516 phenotypes, 513–514 Medulla oblongata, thiamine triphosphatase, 53 Megaloblastic anemia syndrome, thiamine-responsive, 549–564 animal model, 557–560 biochemical aspects, 552–554 clinical manifestations, 550–551 genetics, 554–557 treatment, 551–552

Index Methionine 250, bacterial acetohydroxyacid synthases, 238–239 Methoxythiamine pyrophosphate, conversion from antibiotic bacimethrin, by thiamine pyrophosphate biosynthetic enzymes, 24 3-methyl-2-oxobutanoate dehydrogenase (lipoamide), 94 3-methyl-branched fatty acid metabolism, thiamine pyrophosphate, 525–538 alpha-oxidation, 3-methyl-branched fatty acids, 527–529 alpha-oxidation pathway, 2-hydroxyphytanoyl-COa lyase, thiaminedependent enzyme, 530–534 phytanic acid, deﬁcient breakdown of, 529–530 Microcephaly, with pyruvate dehydrogenase complex deﬁciency, 473 Mitochondrial dysfunction, oxidative stress, neurodegeneration cycle, 544–546 Mitochondrial medium-chain acylCoA synthetase, 348–351 mRNA distribution, thiamine triphosphate, thiamine triphosphatase, human tissue, 53 Multiple-tissue-expression array, human, by Clontech, 53 Nervous system, thiamine triphosphatase, 53 Neuroblastoma cells, thiamine metabolism in, 45, 46 Neuronal loss, in Wernicke–Korsakoﬀ syndrome, oxidative stress, 539–548 mitochondrial dysfunction, oxidative stress and, neurodegeneration cycle, 544–546

579 [Neuronal loss, in Wernicke–Korsakoﬀ syndrome, oxidative stress] thiamine deﬁciency: mitochondrial dysfunction in, 541–542 NMDA receptor-mediated exitotoxicity in, 542–543 thiamine esters, 540–541 thiamine-deﬁciency-related neuronal cell death, oxidative stress and, 543–544 thiamine-dependent enzymes, 540–541 Neuropathy, with pyruvate dehydrogenase complex deﬁciency, 473 Nucleus accumbens, thiamine triphosphatase, 53 Occipital lobe, brain, thiamine triphosphatase, 53 One-electron processes, role of thiamine diphosphate, 11–12 ‘‘Orphan’’ thiamine diphosphate decarboxylases, carbon-carbon bonding reactions, thiaminedependent enzymes as catalysts, 77–93 acyloin condensations, 78 C-1 metabolism, 82–84 imines as acceptors, 84–86 ketones as acceptors, 84–86 range of reactions catalyzed by, 79–82 Ovary, thiamine triphosphatase, 53 Oxalyl-CoA decarboxylase, 94 Oxidative stress, in Wernicke–Korsakoﬀ syndrome neuronal loss, 539–548 mitochondrial dysfunction, oxidative stress and, neurodegeneration cycle, 544–546 thiamine deﬁciency: mitochondrial dysfunction in, 541–542

580 [Oxidative stress, in Wernicke–Korsakoﬀ syndrome neuronal loss] NMDA receptor-mediated exitotoxicity in, 542–543 thiamine esters, 540–541 thiamine-deﬁciency-related neuronal cell death, oxidative stress and, 543–544 thiamine-dependent enzymes, 540–541 2-oxoglutarate decarboxylase, 94 Oxoglutarate dehydrogenase (lipoamide), 94 Pancreas, thiamine triphosphatase, 53 Parietal lobe, brain, thiamine triphosphatase, 53 Peripheral neuropathy, with pyruvate dehydrogenase complex deﬁciency, 473 Phenylglyoxylate dehydrogenase (acylating), 94 Phosphoketolase, 94 Phospho-pyruvate dehydrogenase phosphatase, 485, 487, 492, 496, 502 (See also PDP within text): down-regulation of, 496 isoenzymes of, 486 PDP1, 492 dephosphorylation by, 492 Phosphorylation, in pyruvate dehydrogenase, thiamine pyrophosphate interaction, 485, 486, 492, 496, 498, 500, 502 level of, 496 rates of, 492, 496 rate-speciﬁc, 496 regulation by, 496 sites, 496, 500, 502 site-speciﬁc, 496 three, 492, 496 Placenta, thiamine triphosphatase, 53 Pons, thiamine triphosphatase, 53

Index Potassium phosphate concentration, eﬀect on yeast acetohydroxyacid synthase, 255, 256 Prostate, thiamine triphosphatase, 53 Protein lipoylation, lipoic acid in, 343–362 lipoate-activating enzyme, puriﬁcation, characterization of, 344–348 lipoic acid, activation of, 344–351 lipoyltransferase: cDNA structure, 353–356 properties of, 351–353 mitochondrial medium-chain acylCoA synthetase, lipoate-activating enzyme, 348–351 proteins, transfer of lipoic acid to, 351–356 Putamen, thiamine triphosphatase, 53 Pyrimidine phosphate 7, formation of, 20, 21 Pyrimidine pyrophosphate formation, thiamine biosynthetic enzymes, 20–22 Pyrithiamine binding, thiamine pyrophosphokinase, 39 Pyrithiamine-complexed structure, thiamine pyrophosphokinase, 38–40 Pyruvamide, in presence of, least squares alignment of structures of yeast pyruvate decarboxylase, 178, 179 Pyruvate, ferredoxin oxidoreductase, thiazolium ring structures, in reaction intermediates, 168, 169 Pyruvate decarboxylase, 94, 173–216 (See also Zymomonas mobilis): 1, 4-IMINOTDP tautomer on, evidence for presence of, 199–201 active site of yeast pyruvate decarboxylase, ball-and-stick representation, 179, 180 active-center residues, assignment of function to, 201–209

Index [Pyruvate decarboxylase] alternate activation pathway, beta sheets, loosely associated dimer pairs, breakage of interactions between, 186 alternating active sites in functional dimer, mechanism involving, 191–199 bound intermediates, wild type, 207, 208 carbon kinetic isotope eﬀects, 217– 232 carbon isotope eﬀects, 224 carboxyl side chains in, 220, 222 isotope eﬀects, on kcat for yeast pyruvate decarboxylase variants, 224–226 kinetic isotope eﬀects, transitionstate theory, 218–219 low pyruvate concentrations, for pyruvate decarboxylase variants, isotope eﬀects at, 226–228 solvent isotope eﬀects, 223–224 steady-state kinetics, variants, 220–222 wild-type, rate-limiting steps, 220 complete yeast pyruvate decarboxylase tetramer, 176, 177, 178 crystal data, crystallized in presence, absence of eﬀectors, 185 crystallographic models, 95, 98 dimer pair, tightly associated, 175, 176 E477Q yeast pyruvate decarboxylase variant, circular dichroism spectra, 200, 201 ketomalonate, least squares alignment of structures of yeast pyruvate decarboxylase, 178, 179 reactions catalyzed by, 275, 276 regulatory site, active site, activation pathway between, 183, 184 role in mechanism of reaction of thiamine diphosphate, 2

581 [Pyruvate decarboxylase] second conformation, wild-type, kinetic parameters of, 189 structure, 173, 174–182 active-site environment, 179–182 dimer, 175–176 monomer, 174–175 varying tetrameric assemblies of, 176–179 substrate activation, 183–191 thiamine diphosphate, 61–67 reaction mechanisms, 2 thiamine diphosphate cofactor, potential states of, 181, 182 third conformation, wild-type, kinetic parameters of, 190 wild type, acetaldehyde formation by, progress curves of, 188, 189 wild-type, 190 carbon kinetic isotope eﬀects, 220 Zymomonas mobilis, 63, 190 Pyruvate dehydrogenase, 94, 485–487, 489, 492, 494–496, 500, 502 (See also E1 within text): activation of, 487, 489, 494–496, 500, 502 active site of, 492, 495 alpha-subunit of, 486, 492, 494, 495 Ascaris suum, 496 beta-subunit, 494, 495 binding of, 500 Caenorhabditis elegans, 496 catalysis, 486 deﬁciency, 487, 492 degradation, 494 dephosphorylation of, 485 E1-binding domain, 485 E1-deﬁciency, 486, 487, 494 fruit ﬂy, 496 function, 492, 494, 502 genetic defects of, 487 human, 486, 487, 489, 495, 496 PDC-E1 structure, 486 inactivation of, 486, 495, 496 isoforms of, 487

582 [Pyruvate dehydrogenase] kinetic parameters of, 492 mammalian, 496 multiple sites of, 500 mutant, 492–496, 498, 500, 502 phosphorylated, 496 phosphorylation of, 496 sites of, 500, 502 plant, 496 reaction, 495, 496, 498 reactivation, 492 regulation of, 486, 496 site-directed mutants of, 486 speciﬁc activity, 496 structure-function of, 487 thiamine diphosphate, 71 thiamine pyrophosphate, interactions: inhibitor: site-directed, 495 tight-binding, 489 kinetic analysis, 486, 495, 496, 502 lag-phase, 487, 489, 495 mutagenesis, 494 site-directed, 486, 495, 496 mutation, 487, 489, 492–495, 502 (See also Mutations within text): missense, 487 patient’s, 493 point, 487 recreating, 487 substitution, 486 wild-type, 495, 500 yeast, 496 Pyruvate dehydrogenase complex, 309–330, 331–342, 485, 486, 494, 500 (See also PDC within text): active, 496 active-site coupling, 333–334 activity, 486, 487, 492, 494, 496, 500, 502 assay, 495, 496, 498 assembly, 494 of Bacillus stearothermophilus, 332, 333

Index [Pyruvate dehydrogenase complex] bovine kidney PDHC, atomic structures, 323, 324–325 catalysis, 339–340 components, 487, 496 computer image processing, 313–314 conformational variability, of ‘‘breathing’’ E2 core, 317–321 cryoEM, 314–315 deﬁciency, 471–485, 487, 492 clinical, 472–474 features, frequency of, 473 genetic basis, 474–475 metabolic consequences, 472–474 outcome, 475–476 pathological consequences, 472–474 treatment, 476–479 electron density map, E1E2 subcomplex, 337, 338 electron microscopy, 313 Escherichia coli replacement by: aceE-IpdA region, restructuring, operon, 393–395 changing coenzyme speciﬁcities, PDH, ODH complexes, 389–393 genetically engineered variants, 387–406 lipoyl domains, per acetyl transferase chain, 395–399 pyruvate oxidase, replacing PDH complex by, physiological consequences of, 399–401 Escherichia coli replacement by pyruvate oxidase, 387–406 lipoly-lysine residue, lipoyl domain, inserted into active site of E1 component, 334, 335 lipoyl domain, 334, 335 interaction with E1, 334–337 reaction mechanism, 334–337 lipoyl domain-mediated activated function, control, PD kinases, phosphatase 1, 363–386 composition, 363–364

Index [Pyruvate dehydrogenase complex] E2-domain structures, domain roles, 366–368 E2-facilitated kinase function, 370–377 E3BP-domain structures, domain roles, 366–368 kinase activity, E2-mediated, modiﬁed regulation of, 373–377 organization of, 368–370 PDK2, binding, activation of, 370–372 PDK3, binding, activation of, 372–373 PDK4, binding speciﬁcity, regulation, 377 PDP1, E2-mediated CA2+ activation of, 377–379 regulatory enzymes, 364–365 roles, 363–364 localization of E2 in tE2/BP/E3 complex, size variation classiﬁcation, 321–323 mammalian PDHC, reactions catalyzed by, 309, 310 molecular architecture, 337–339 native: preparation of, 312–313 recombinant subcomplexes, 312–313 particle size variation, assessment of, 314 reaction, 489, 495 recombinant S. cerevisiae tE2 core, size distribution, 317, 318–319 reconstitution in, 487, 492 reconstruction, computer image, 313–314 regulation, 496 schematic reaction mechanism, 336, 337 size, of ‘‘breathing’’ E2 core, 317–321 substrate channeling, 333–334

583 [Pyruvate dehydrogenase complex] three-dimensional structures: conformational variability, of ‘‘breathing’’ E2 core, 317–321 localization of E2 in tE2/BP/E3 complex, size variation classiﬁcation, 321–323 size, of ‘‘breathing’’ E2 core, 317–321 yeast tE2 core, tE2 complexed with binding protein, 315–316 three-dimensional structures, 315–316 Pyruvate dehydrogenase complex kinase 2, phosphatase 1, binding, activation of, 370–372 Pyruvate dehydrogenase complex kinase 3, binding, activation of, 372–373 Pyruvate dehydrogenase complex kinase 4, binding speciﬁcity, regulation, 377 Pyruvate dehydrogenase kinase, 433–448, 485, 492, 496, 502 (See also PDK within text): activity of, 496, 502 binding of, 502 bovine, 496 catalytic mechanism, 440–442 distribution of PDK isozymes, 500 isozymes of, 486, 492, 500, 502 nucleotide binding mechanism, 437–440 PDK1, 492, 500 PDK2, 492, 500, 502 PDK3, 500, 502 PDK4, 500, 502 regulation of, by altered expression of kinase, 442–445 three-dimensional structure, 435–437 up-regulation of, 496, 500 Pyruvate oxidase, 94 covalent intermediates, 68, 69

584 [Pyruvate oxidase] recombinant wild type, stabilized mutant, 95, 96 thiamine diphosphate, 68–71 Radioactivities, thiamine derivatives, 45 Rectum, thiamine triphosphatase, 53 Reproductive tract, thiamine triphosphatase, 53 Saccharomyces cerevisiae, transketolase, recombinant wild type, 95, 97 Salivary gland, thiamine triphosphatase, 53 Seizure disorder, with pyruvate dehydrogenase complex deﬁciency, 473 Ser26, benzoylformate decarboxylase, 139–140 Single-gene disorders, complexity of, maple syrup urine disease, thiamine responsiveness, 509–524 Skeletal muscle, thiamine triphosphatase, 53 Spleen, fetal, thiamine triphosphatase, 53 Stomach, thiamine triphosphatase, 53 Tartronate-semialdehyde synthase, 94 Temporal lobe, brain, thiamine triphosphatase, 53 Testis, thiamine triphosphatase, 53 Thalamus, thiamine triphosphatase, 53 Thiaminase, X-ray structure of, 24 Thiaminase I, mechanistic proposal for, 24, 25 Thiamine, 493, 494 biosynthetic enzymes, 15–28 antibiotic bacimethrin, conversion to methoxythiamine pyrophosphate, 24 bacterial thiamine biosynthetic pathway, 15, 16

Index [Thiamine] HMP-P kinase, X-ray structure of, 20, 22 NifS, partial reactions catalyzed by, identiﬁcation, 15, 17 pyrimidine phosphate 7, formation of, 20, 21 thiaminase, X-ray structure of, 24 thiaminase I, mechanistic proposal for, 24, 25 thiamine phosphate synthase, X-ray structure of, 22 thiamine pyrophosphate, enzymatic synthesis of, 23, 24 thiazole kinase, X-ray structure of, 19 thiazole phosphate 4, formation of, 16, 17 ThiF, partial reactions catalyzed by, identiﬁcation, 15, 17 ThiG, partial reactions catalyzed by, identiﬁcation, 15, 17 ThiO: partial reactions catalyzed by, identiﬁcation, 15, 17 X-ray structure of, 18, 19 ThiS, partial reactions catalyzed by, identiﬁcation, 15, 17 pyrophosphate, 486, 487, 489, 495, 500 (See also TPP within text): aﬃnity for, 489, 492, 495 aminopyrimidine ring of, 486, 495 binding, 492, 494, 495, 500, 502 region, 486, 500 site, 492, 494, 495, 502 dependent enzymes, 486, 495 Km for, 487, 492–495, 500, 502 motif, 486, 487, 489, 492 pyrophosphate moiety of, 486, 487, 492 requiring enzymes, 486, 487 thiazolium ring of, 486 thiothiazolone pyrophosphate, 489 (See also TTTPP within text) treatment, 492, 493

Index Thiamine biosynthetic enzymes: bacimethrin toxicity, 23–24 enzymatic synthesis, thiamine pyrophosphate, 23 mechanistic studies, 15–28 pyrimidine pyrophosphate formation, 20–22 structural studies, 15–28 thiamine degradation, 24–26 thiamine pyrophosphate formation, 22–23 thiazole phosphate formation, 15–20 Thiamine deﬁciency: mitochondrial dysfunction in, 541–542 NMDA receptor-mediated exitotoxicity in, 542–543 oxidative stress, neuronal cell death, 543–544 Thiamine degradation, thiamine biosynthetic enzymes, 24–26 Thiamine diphosphate: acetohydroxyacid synthase, 264 chemical intermediates in catalysis by, 1–14 acetyl-thiamine diphosphate, in aqueous solutions, structures of, 9, 10 acyl-thiamine diphosphates, as intermediates, 4–11 enzymatic intermediate, identiﬁcation of as, 9–11 ﬂuoropyruvate, as alternative substrate leading to, 7–8 in reactions of alpha-ketoacid dehydrogenase complexes, 5–7 synthesis of, 8–9 aldehyde addition compounds with, 4 alpha-ketoacid dehydrogenase complexes, reaction mechanisms of, 6 enamine-thiamine diphosphate and acyl-thiamine diphosphate intermediates, role of, 3, 5

585 [Thiamine diphosphate] ﬂuoropyruvate, pyruvate dihydrogenase complex, substrate for E1 component of, 8 free-radical reactions, role in, 11–12 one-electron processes, role in, 11–12 pyruvate decarboxylase, role in mechanism of reaction of, 2 Thiamine diphosphate-dependent enzymes, 57–76 acetohydroxyacid synthase, 67–68 comparison with, 261–263 covalent intermediates, 68, 69 catalysis, model of, 66 covalent intermediates, 68, 69 in enzymic thiamine catalysis, 61–72 in thiamine diphosphate catalysis, 58 ligand-induced conformational changes, 93–112 crystal, solution structures, compared, 93–112 Lactobacillus plantarum, 95, 96 ligand-induced conformational changes, pyruvate oxidase, 99–102 pyruvate decarboxylases, crystallographic models, 95, 98 recombinant wild type, pyruvate oxidase, stabilized mutant, 95, 96 Saccharomyces cerevisiae, transketolase from, recombinant wild type, 95, 97 pyruvate decarboxylase, 61–67 pyruvate dehydrogenase, 71 pyruvate oxidase, 68–71 covalent intermediates, 68, 69 superfamily of thiamine diphosphate-dependent enzymes, covalent intermediates, 68, 69 thiamine, 57–76

586 [Thiamine diphosphate-dependent enzymes] acetohydroxyacid synthases, 67–68 covalent intermediates, in enzymic thiamine catalysis, 61–72 pyruvate decarboxylase, 61–67 pyruvate dehydrogenase, 71 pyruvate oxidase, 68–71 transketolase, 71–72 transketolase, 71–72 covalent intermediates, 68, 69 Zymomonas mobilis, pyruvate decarboxylase, active site of, 63 Thiamine phosphate synthase, X-ray structure of, 22 Thiamine pyrophosphate: 3-methyl-branched fatty acid metabolism, 525–538 alpha-oxidation, 3-methylbranched fatty acids, 527–529 alpha-oxidation pathway, 2hydroxyphytanoyl-COa lyase, thiamine-dependent enzyme, 530–534 phytanic acid, deﬁcient breakdown of, 529–530 enzymatic synthesis, 23, 24 thiamine biosynthetic enzymes, 23 formation, thiamine biosynthetic enzymes, 22–23 structure, thiamine pyrophosphokinase, 34–38 Thiamine pyrophosphokinase, 29–42 2Fo-Fc maps, 34, 35 binding in mouse thiamine pyrophosphokinase, 34, 36 buﬀers, activity on, 31, 32 crystal soaking, 30 crystallization, 30 crystallographic data, 31 ethanol, eﬀect on mouse thiamine pyrophosphokinase activity, 32, 33 expression, 30

Index [Thiamine pyrophosphokinase] inhibitors of, 34 kinetic studies, 31–34 kinetics, 30 pyrithiamine binding in, 39 pyrithiamine-complexed structure, 38–40 reﬁnement, 30–31 statistics, 31 structure determination, 30–31 substrates, 34 superposition of from mouse thiamine pyrophosphokinase, 37 thiamine pyrophosphate structure, 34–38 thiamine ring atoms, numerical assignment of, 38 Thiamine responsiveness, maple syrup urine disease, 509–524 branched-chain alpha-ketoacid dehydrogenase, 510 regulation of, 510–512 by complex-speciﬁc kinase, 512–513 role of thiamine pyrophosphate in, 517–518 history of disease, 509–510 model systems, 518–519 mutations, 514–516 phenotypes, 513–514 thiamine-responsive, 516–517 Thiamine ring atoms, numerical assignment of, 38 Thiamine triphosphatase, 43–56 24-KDA, 52 adenylate kinase activities in tissue, 47 cellular role of, 43–56 molecular characterization of, 47–51 mRNA distribution, human tissue, 53 Thiamine triphosphate, 43–56 adenylate kinase activities in tissue, 47

Index [Thiamine triphosphate] cellular role of, 43–56 mRNA distribution, human tissue, 53 protein phosphorylation, 44–45 synthesis, mechanism of, 46–47 turnover, 45–46 Thiamine-responsive megaloblastic anemia syndrome, 549–564 animal model, 557–560 biochemical aspects, 552–554 clinical manifestations, 550–551 genetics, 554–557 treatment, 551–552 Thiazole kinase, X-ray structure of, 19 Thiazole phosphate 4, formation of, 16, 17 Thiazole phosphate formation, thiamine biosynthetic enzymes, 15– 20 Thiazolium ring structures, ferredoxin oxidoreductase, in reaction intermediates, 168, 169 Thymus, fetal, thiamine triphosphatase, 53 Thyroid gland, thiamine triphosphatase, 53 Trachea, thiamine triphosphatase, 53 Transketolase, 94 alpha-carbanion/enamine reaction intermediate in kinetic crystallography, 159–172 beta hydroxylpyruvic acid binding to transketolase, circular dichroism measurement, kinetics of, 162–163 circular dichroism measurements, 161 conformational changes, intermediate formation and, 167–168 crystallographic model building, reﬁnement, 162 crystallography, 161 data collection, 161

587 [Transketolase] dihydroxy ethyl-thiamine diphosphate, interactions with activesite residues, 165–167 dihydroxy ethyl-thiamine diphosphate intermediate, 164–165 near-UV CD spectra reﬂecting formation of, 167 election density maps, 163 protein expression, puriﬁcation, 161 transketolase-beta hydroxylpyruvic acid adduct, kinetics of formation of, 161 covalent intermediates, 68, 69 ferredoxin oxidoreductase, alphacarbanion/enamine reaction intermediate in transketolase site, 168, 169 thiamine diphosphate, 71–72 Tris-HC1 phosphate, mouse thiamine pyrophosphokinase, activity on, 31, 32 Trp464, properties of, bacterial acetohydroxyacid synthases, 237 Ulfoacetaldehyde lyase, 94 Urinary tract, thiamine triphosphatase, 53 Uterus, thiamine triphosphatase, 53 Val375, properties of, bacterial acetohydroxyacid synthases, 237 Val461a, properties of, bacterial acetohydroxyacid synthases, 237 Ventricle, thiamine triphosphatase, 53 Wernicke-Korsakoﬀ syndrome, neuronal loss in, oxidative stress, 539– 548 mitochondrial dysfunction, oxidative stress and, neurodegeneration cycle, 544–546

588 [Wernicke-Korsakoﬀ syndrome, neuronal loss in, oxidative stress] thiamine deﬁciency: mitochondrial dysfunction in, 541–542 neuronal cell death, oxidative stress and, 543–544 NMDA receptor-mediated exitotoxicity in, 542–543 thiamine esters, 540–541 thiamine-dependent enzymes, 540–541

Index Yeast: acetohydroxyacid synthase (See Acetohydroxyacid synthase) pyruvate decarboxylase (See Pyruvate decarboxylase) thiamine triphosphatase (See Thiamine triphosphatase)

Zymomonas mobilis, pyruvate decarboxylase, 63, 190