Metal-Carbon Bonds in Enzymes and Cofactors

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METAL IONS IN LIFE SCIENCES VOLUME 6

Metal-Carbon Bonds in Enzymes and Cofactors

METAL IONS IN LIFE SCIENCES edited by Astrid Sigel,(1) Helmut Sigel,(1) and Roland K. O. Sigel(2) (1)

(2)

Department of Chemistry Inorganic Chemistry University of Basel Spitalstrasse 51 CH-4056 Basel, Switzerland Institute of Inorganic Chemistry University of Zu¨rich Winterthurerstrasse 190 CH-8057 Zu¨rich, Switzerland

VOLUME 6

Metal-Carbon Bonds in Enzymes and Cofactors

The figure on the dust cover shows coenzyme B12 (¼ 5 0 -deoxy-5 0 adenosylcobalamin) containing a cobalt-carbon bond; prepared by Roland K. O. Sigel using the CSD coordinates DADCBL.

ISBN: 978-1-84755-915-9 ISSN: 1559-0836 DOI: 10.1039/9781847559159 A catalogue record for this book is available from the British Library r Royal Society of Chemistry 2009 All rights reserved Apart from fair dealing for the purposes of research for non-commercial purposes or for private study, criticism or review, as permitted under the Copyright, Designs and Patents Act 1988 and the Copyright and Related Rights Regulations 2003, this publication may not be reproduced, stored or transmitted, in any form or by any means, without the prior permission in writing of The Royal Society of Chemistry or the copyright owner, or in the case of reproduction in accordance with the terms of licences issued by the Copyright Licensing Agency in the UK, or in accordance with the terms of the licences issued by the appropriate Reproduction Rights Organization outside the UK. Enquiries concerning reproduction outside the terms stated here should be sent to The Royal Society of Chemistry at the address printed on this page. Published by The Royal Society of Chemistry, Thomas Graham House, Science Park, Milton Road, Cambridge CB4 0WF, UK Registered Charity Number 207890 For further information see our web site at www.rsc.org

Historical Development and Perspectives of the Series Metal Ions in Life Sciences

It is an old wisdom that metals are indispensable for life. Indeed, several of them, like sodium, potassium, and calcium, are easily discovered in living matter. However, the role of metals and their impact on life remained largely hidden until inorganic chemistry and coordination chemistry experienced a pronounced revival in the 1950s. The experimental and theoretical tools created in this period and their application to biochemical problems led to the development of the field or discipline now known as Bioinorganic Chemistry, Inorganic Biochemistry, or more recently also often addressed as Biological Inorganic Chemistry. By 1970 Bioinorganic Chemistry was established and further promoted by the book series Metal Ions in Biological Systems founded in 1973 (edited by H.S., who was soon joined by A.S.) and published by Marcel Dekker, Inc., New York, for more than 30 years. After this company ceased to be a family endeavor and its acquisition by another company, we decided, after having edited 44 volumes of the MIBS series (the last two together with R.K.O.S.) to launch a new and broader minded series to cover today’s needs in the Life Sciences. Therefore, the Sigels new series is entitled Metal Ions in Life Sciences. After publication of the first four volumes (2006–2008) with John Wiley & Sons, Ltd., Chichester, UK, we are happy to join forces now in this still new endeavor with the Royal Society of Chemistry, Cambridge, UK; a most experienced Publisher in the Sciences.

*

Reproduced with some alterations by permission of John Wiley & Sons, Ltd., Chichester, UK (copyright 2006) from pages v and vi of Volume 1 of the series Metal Ions in Life Sciences (MILS-1).

vi

PERSPECTIVES OF THE SERIES

The development of Biological Inorganic Chemistry during the past 40 years was and still is driven by several factors; among these are (i) the attempts to reveal the interplay between metal ions and peptides, nucleotides, hormones or vitamins, etc., (ii) the efforts regarding the understanding of accumulation, transport, metabolism and toxicity of metal ions, (iii) the development and application of metal-based drugs, (iv) biomimetic syntheses with the aim to understand biological processes as well as to create efficient catalysts, (v) the determination of high-resolution structures of proteins, nucleic acids, and other biomolecules, (vi) the utilization of powerful spectroscopic tools allowing studies of structures and dynamics, and (vii), more recently, the widespread use of macromolecular engineering to create new biologically relevant structures at will. All this and more is and will be reflected in the volumes of the series Metal Ions in Life Sciences. The importance of metal ions to the vital functions of living organisms, hence, to their health and well-being, is nowadays well accepted. However, in spite of all the progress made, we are still only at the brink of understanding these processes. Therefore, the series Metal Ions in Life Sciences will endeavor to link coordination chemistry and biochemistry in their widest sense. Despite the evident expectation that a great deal of future outstanding discoveries will be made in the interdisciplinary areas of science, there are still ‘‘language’’ barriers between the historically separate spheres of chemistry, biology, medicine, and physics. Thus, it is one of the aims of this series to catalyze mutual ‘‘understanding’’. It is our hope that Metal Ions in Life Sciences proves a stimulus for new activities in the fascinating ‘‘field’’ of Biological Inorganic Chemistry. If so, it will well serve its purpose and be a rewarding result for the efforts spent by the authors. Astrid Sigel, Helmut Sigel Department of Chemistry Inorganic Chemistry University of Basel CH-4056 Basel Switzerland

Roland K. O. Sigel Institute of Inorganic Chemistry University of Zu¨rich CH-8057 Zu¨rich Switzerland October 2005 and October 2008

Preface to Volume 6 Metal-Carbon Bonds in Enzymes and Cofactors

This is the 6th volume within the MILS series; together with the 44 volumes published in our former series Metal Ions in Biological Systems this sums up to in total 50 books. This event is celebrated with a comprehensive Author Index, given at the end of this book. It encompasses the names of all colleagues who contributed to these 44 MIBS and 6 MILS volumes. All these authors deserve our special thanks; without their excellent contributions the two series could not have been successful. The present Volume 6 is devoted to naturally occurring metal-carbon bonds, a topic recently obtaining (again) significant momentum, largely – but not only – due to new insights gained with hydrogenases. The field started out about 50 years ago when coenzyme B12 was identified as organometallic derivative of vitamin B12. This moved the cobalt-carbon bond into the center of interest and consequently, the first two chapters of this book are devoted to the organometallic chemistry of B12 coenzymes and to the biochemistry of cobalamin- and corrinoid-dependent enzymes. B12 coenzymes are required in the metabolism of a broad range of organisms including humans; however, only microorganisms have the ability to biosynthesize B12 and other natural corrinoids. This fact alone, together with new metabolic insights (e.g., riboswitches), guarantees a continued fascination – not only for the B12 community. Related to Co-corrin, the Ni-porphinoid unit (F430) is the prosthetic group of methyl-coenzyme M reductase. This enzyme, the topic of Chapter 3, catalyzes the methane-forming step in methanogenic archaea and most

Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-FP007

viii

PREFACE TO VOLUME 6

probably also the methane-oxidizing step in methanotrophic archaea. Chapter 4 deals with acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, i.e., bifunctional nickel-containing enzymes, which catalyze the synthesis of acetyl-CoA and the reversible reduction of CO2 to CO in anaerobic, mostly thermophilic, organisms, able to grow chemiautotrophically on simple inorganic compounds like CO2. Ni-C bonds with methyl, acetyl, carbonyl, and carboxylate groups are evidenced. [NiFe]-, [FeFe]-, and [Fe]-hydrogenases are detailed in the next three chapters. These enzymes, present in many microorganisms, catalyze the oxidation of molecular hydrogen or the reduction of protons. All of them have a Fe(CO)x unit in their active site. Iron-cyanide units occur in [NiFe]and [FeFe]-hydrogenases. However, despite the indicated similarities they clearly have independent evolutionary origins. The participation of the commonly considered toxic ligands CO and CN– in the active sites of hydrogenases is still a surprise to many; yet, exactly their occurrence incites a great interest in physical chemists as well as evolutionary biologists. The dual role of heme as cofactor and substrate in the biosynthesis of carbon monoxide is the topic of Chapter 8. Carbon monoxide is a ubiquitous molecule in the atmosphere but it is also produced in mammalian, plastidic, and bacterial cells as a byproduct in the catalytic cycle of heme degradation as catalyzed by the enzyme heme oxygenase. Most fascinating is the fact that the biological role of CO spans the range from toxic to cytoprotective, depending on its concentration. CO generated by heme oxygenase is now known to function in several important physiological processes, including vasodilation, apoptosis, inflammation, and possibly neurotransmission. The relevance of the copper-carbon bond in biological inorganic chemistry will probably not easily come to the mind of most biochemical and inorganic researchers. However, there is a vast amount of literature, cunningly presented in Chapter 9. CO as well as CN– have proven very useful in obtaining insights into the active site structures and mechanisms of copper proteins. Naturally, in these instances both ligands are inhibitors and used as probes. However, there is also the recently described copper-carbon unit present in a carbon monoxide dehydrogenase, which contains a novel molybdenum-copper catalytic site, or the copper(I)-arene unit, which was evidenced in a bacterial copper chaperone. Apparently also a plant receptor site (ETR1) utilizes Cu(I) to sense the growth hormone ethylene. Chapter 10 focuses on the interaction of CN– with enzymes containing vanadium, manganese, non-heme iron, and zinc, and the inhibiting properties of this ligand, allowing its use as a probe. The reaction mechanism of the molybdenum hydroxylase xanthine oxidoreductase is revisited in Chapter 11; previously a molybdenum-carbon bond was postulated but now proof is presented against its formation. The terminating Chapter 12 reviews

PREFACE TO VOLUME 6

ix

briefly the most popular computational approaches employed in theoretical studies of bioorganometallic species by providing detailed examples. Taken together, MILS-6 summarizes our knowledge on Metal-Carbon Bonds in Enzymes and Cofactors; i.e., it emphasizes the role of metal-carbon bonds for life as well as research. However, there are many metal-carbon bonds which occur in the environment in compounds like alkyl-arsenicals or -mercurials and in lead- or tinorganyls, etc., most of them known as being toxic. Consequently, the next volume (MILS-7) will be devoted to Organometallics in Environment and Toxicology. Astrid Sigel Helmut Sigel Roland K. O. Sigel

Contents

HISTORICAL DEVELOPMENT AND PERSPECTIVES OF THE SERIES PREFACE TO VOLUME 6

v vii

CONTRIBUTORS TO VOLUME 6

xvii

TITLES OF VOLUMES 1–44 IN THE METAL IONS IN BIOLOGICAL SYSTEMS SERIES

xix

CONTENTS OF VOLUMES IN THE METAL IONS IN LIFE SCIENCES SERIES

xxi

1

ORGANOMETALLIC CHEMISTRY OF B12 COENZYMES Bernhard Kraütler

1

Abstract 1. Introduction 2. Structure of B12 Derivatives in the Crystal and in Solution 3. Redox Chemistry of B12 Derivatives 4. Reactivity of B12 Derivatives in Organometallic Reactions 5. Organometallic B12 Derivatives as Cofactors and Intermediates in Enzymes 6. Concluding Remarks and Future Directions Acknowledgments Abbreviations References

2 2 5 18 24

Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-FP011

34 40 41 41 42

xii

2

CONTENTS

COBALAMIN- AND CORRINOID-DEPENDENT ENZYMES Rowena G. Matthews Abstract 1. Introduction. What Is a Corrinoid? 2. Corrinoid-Dependent Methyltransferases 3. Adenosylcobalamin-Dependent Rearrangements and Eliminations 4. Concluding Remarks Acknowledgments Abbreviations and Definitions References

3

NICKEL-ALKYL BOND FORMATION IN THE ACTIVE SITE OF METHYL-COENZYME M REDUCTASE Bernhard Jaun and Rudolf K. Thauer Abstract 1. Introduction 2. Nickel-Carbon Bond Formation in Free Coenzyme F430 3. Nickel-Alkyl Bond Formation in MCR Upon Inactivation with Alkyl Halides 4. Methyl-Nickel Bond Formation in Methyl-Coenzyme M Reductase During Catalysis? 5. Observations to Be Followed Up Acknowledgments Abbreviations References

4

NICKEL-CARBON BONDS IN ACETYL-COENZYME A SYNTHASES/CARBON MONOXIDE DEHYDROGENASES Paul A. Lindahl Abstract 1. Introduction 2. Redox and Catalytic Properties of the A- and C-Clusters 3. Evidence for a Ni-CO Bond in the Ared-CO State of the A-cluster

53 54 54 56 84 106 107 107 107

115 116 116 119 120 123 129 129 129 130

133 133 134 137 139

CONTENTS

Evidence for a Ni-CH3 Bond in the Methylated Intermediate of the A-Cluster 5. Evidence for a Ni-C(O)CH3 Bond in the Acetyl Intermediate of the A-Cluster 6. Evidence for a Ni-CO Bond in the C-Cluster 7. Evidence for a Ni-C(O)O-Fe Bond in the C-Cluster 8. Conclusions and Future Studies Acknowledgment Abbreviations and Definitions References

xiii

4.

5

6

139 140 141 143 144 147 147 147

STRUCTURE AND FUNCTION OF [NiFe]-HYDROGENASES Juan C. Fontecilla-Camps

151

Abstract 1. Introduction 2. Hydrogenase Structure 3. Hydrogenase Maturation and Active Site Assembly 4. Electron Transfer 5. Proton Transfer 6. Oxidized Inactive States of the [NiFe]-Hydrogenase Active Site 7. Substrate Binding and Catalysis 8. Concluding Remarks Acknowledgments Abbreviations References

152 152 153 160 165 166 168 172 173 173 173 174

CARBON MONOXIDE AND CYANIDE LIGANDS IN THE ACTIVE SITE OF [FeFe]-HYDROGENASES John W. Peters

179

Abstract 1. Introduction 2. [FeFe]-Hydrogenase Structure 3. [FeFe]-Hydrogenase Spectroscopic Studies 4. H-Cluster Model Complexes 5. H-Cluster Biosynthesis 6. Future Directions Acknowledgments

180 180 181 192 195 199 206 207

xiv

7

CONTENTS

Abbreviations and Definitions References

208 208

CARBON MONOXIDE AS INTRINSIC LIGAND TO IRON IN THE ACTIVE SITE OF [Fe]-HYDROGENASE Seigo Shima, Rudolf K. Thauer, and Ulrich Ermler

219

Abstract 1. Introduction 2. Physiology 3. The Iron Guanylylpyridinol Cofactor in the Enzyme-Free State 4. Structure of [Fe]-Hydrogenase with and without the Iron Guanylylpyridinol Cofactor Bound 5. Ligands to Iron in the Active Site of [Fe]-Hydrogenase 6. Proposed Catalytic Mechanisms 7. Concluding Remarks Acknowledgments Abbreviations References

8

THE DUAL ROLE OF HEME AS COFACTOR AND SUBSTRATE IN THE BIOSYNTHESIS OF CARBON MONOXIDE Mario Rivera and Juan C. Rodrı´guez Abstract 1. Introduction 2. The Biosynthesis of Carbon Monoxide 3. Heme Oxygenase Favors Heme Hydroxylation over Ferryl Formation. The Nature of the Ferric Hydroperoxide Complex in Heme Oxygenase 4. Heme Oxygenase Dynamics and Heme Breakdown. The Distal Ligand Has a Profound Effect in the Dynamic Behavior of Heme Oxygenase 5. The Regioselectivity of Heme Hydroxylation 6. Conclusion and Outlook Acknowledgments Abbreviations References

220 220 222 223 227 231 235 237 237 237 238

241

242 243 247

259

268 276 284 285 286 286

CONTENTS

9

COPPER-CARBON BONDS IN MECHANISTIC AND STRUCTURAL PROBING OF PROTEINS AS WELL AS IN SITUATIONS WHERE COPPER IS A CATALYTIC OR RECEPTOR SITE Heather R. Lucas and Kenneth D. Karlin Abstract 1. Introduction 2. Binuclear Copper Proteins 3. Heterobimetallic Copper-Containing Enzymes 4. Non-Blue Copper Oxidases 5. Blue, Green, and Purple Copper Proteins 6. Copper(I) Recognition Sites or Receptors 7. Miscellaneous 8. General Conclusions Acknowledgments Abbreviations References

10

INTERACTION OF CYANIDE WITH ENZYMES CONTAINING VANADIUM, MANGANESE, NON-HEME IRON, AND ZINC Martha E. Sosa-Torres and Peter M. H. Kroneck Abstract 1. Introduction 2. Vanadium Enzymes 3. Manganese Enzymes 4. Non-Heme Iron Enzymes 5. Zinc Enzymes 6. Conclusions Acknowledgments Abbreviations References

11

THE REACTION MECHANISM OF THE MOLYBDENUM HYDROXYLASE XANTHINE OXIDOREDUCTASE: EVIDENCE AGAINST THE FORMATION OF INTERMEDIATES HAVING METAL-CARBON BONDS Russ Hille Abstract 1. Introduction

xv

295

296 297 298 317 334 337 344 346 349 350 351 352

363

364 364 368 373 377 383 388 388 388 389

395

396 396

xvi

CONTENTS

2.

Electron-Nuclear Double Resonance Studies of the ‘‘Very Rapid’’ Species 3. X-Ray Crystal Structures Relevant to The Reaction Mechanism 4. General Conclusions Acknowledgments Abbreviations and Definitions References

12

400 404 413 414 414 414

COMPUTATIONAL STUDIES OF BIOORGANOMETALLIC ENZYMES AND COFACTORS 417 Matthew D. Liptak, Katherine M. Van Heuvelen, and Thomas C. Brunold Abstract 1. Introduction 2. Computational Approaches to Bioorganometallic Chemistry 3. Formation and Cleavage of the Co–C Bond of Cobalamin in Enzyme Active Sites 4. Organometallic Chemistry and Catalytic Cycle of Methyl-Coenzyme M Reductase 5. Geometric and Electronic Structures of the Carbon Monoxide Dehydrogenase/Acetyl-Coenzyme A Synthase Active Site Clusters 6. Magnetic Properties of the Active Site Cluster of Iron-Only Hydrogenases 7. Concluding Remarks and Future Directions Acknowledgments Abbreviations References

418 419 420 426 435

442 447 450 452 452 454

SUBJECT INDEX

461

AUTHOR INDEX OF CONTRIBUTORS TO MIBS-1 TO MIBS-44 AND MILS-1 TO MILS-6

497

Contributors to Volume 6

Numbers in parentheses indicate the pages on which the authors’ contributions begin. Thomas C. Brunold Department of Chemistry, University of WisconsinMadison, 1101 University Avenue, Madison, WI 53706, USA (Fax: +1-608-262-6143) [email protected] (417) Ulrich Ermler Max Planck Institute for Biophysics, Max-von-Laue-Strasse 3, D-60438 Frankfurt/Main, Germany (219) Juan C. Fontecilla-Camps Laboratoire de Cristallographie et de Cristallogene`se des Proteínes, Institut de Biologie Structurale ‘‘Jean-Pierre Ebel’’ (CEA-CNRS-UJF), 41 rue Jules Horowitz, F-38027 Grenoble Ce´dex 1, France (Fax :+33-4-3878-5122) [email protected] (151) Russ Hille Department of Biochemistry, University of California, Riverside, CA 92521, USA (Fax: +1-951-827-3719) [email protected] (395) Bernhard Jaun Organic Chemistry, ETHZ, ETH Ho¨nggerberg, HCI E317, CH-8093 Zu¨rich, Switzerland [email protected] (115) Kenneth D. Karlin Department of Chemistry, The Johns Hopkins University, New Chemistry 213, 3400 N. Charles Street, Baltimore, MD 21218, USA (Fax: +1-410-516-7044) [email protected] (295) Bernhard Kraütler Institute of Organic Chemistry and Centre of Molecular Biosciences, University of Innsbruck, A-6020 Innsbruck, Austria (Fax: +43-512-507-2892) [email protected] (1) Peter M. H. Kroneck Fachbereich Biologie, Universita¨t Konstanz, Postfach M665, D-78457 Konstanz, Germany [email protected] (363)

xviii

CONTRIBUTORS TO VOLUME 6

Paul A. Lindahl Department of Chemistry and of Biochemistry and Biophysics, Texas A&M University, College Station, TX 77843-3255, USA [email protected] (133) Matthew D. Liptak Department of Chemistry, University of WisconsinMadison, Madison, WI 53706, USA (417) Heather R. Lucas Department of Chemistry, The Johns Hopkins University, New Chemistry 213, 3400 N. Charles Street, Baltimore, MD 21218, USA (Fax: +1-410-516-7044) [email protected] (295) Rowena G. Matthews Department of Biological Chemistry and Life Sciences Institute, University of Michigan, Room 4002, 210 Washtenaw Avenue, Ann Arbor, MI 48109-2216, USA (Fax: +1-734-763-6492) [email protected] (53) John W. Peters Montana State University, Department of Chemistry and Biochemistry, Bozeman, MT 59717, USA (Fax: +1-406-994-7212) [email protected] (179) Mario Rivera Ralph N. Adams Institute for Bioanalytical Chemistry, Department of Chemistry, The University of Kansas, Multidisciplinary Research Building, 2030 Becker Dr., Lawrence, KS 66047, USA (Fax: +1-785-864-5396) [email protected] (241) Juan C. Rodrı´ guez Ralph N. Adams Institute for Bioanalytical Chemistry, Department of Chemistry, The University of Kansas, Multidisciplinary Research Building, 2030 Becker Dr., Lawrence, KS 66047, USA (241) Seigo Shima Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Strasse, D-35043 Marburg, Germany [email protected] (219) Martha E. Sosa-Torres Facultad de Quı´ mica, Universidad Nacional Autońoma de Me´xico, Ciudad Universitaria, Coyoacań, 04510, D.F. Me´xico, Me´xico (Fax: +52-55-5616-2010) [email protected] (363) Rudolf K. Thauer Max Planck Institute for Terrestrial Microbiology, Karlvon-Frisch-Strasse, D-35043 Marburg, Germany (Fax: +49-6421-178-109) [email protected] (115, 219) Katherine M. Van Heuvelen Department of Chemistry, University of Wisconsin-Madison, Madison, WI 53706, USA (417)

Titles of Volumes 1–44 in the Metal Ions in Biological Systems Series edited by the SIGELs and published by Dekker/Taylor & Francis (1973–2005)

Volume 1: Volume 2: Volume 3: Volume 4: Volume 5: Volume 6: Volume 7: Volume 8: Volume 9: Volume 10: Volume 11: Volume 12: Volume 13: Volume 14: Volume 15: Volume 16: Volume Volume Volume Volume Volume

17: 18: 19: 20: 21:

Volume 22: Volume 23:

Simple Complexes Mixed-Ligand Complexes High Molecular Complexes Metal Ions as Probes Reactivity of Coordination Compounds Biological Action of Metal Ions Iron in Model and Natural Compounds Nucleotides and Derivatives: Their Ligating Ambivalency Amino Acids and Derivatives as Ambivalent Ligands Carcinogenicity and Metal Ions Metal Complexes as Anticancer Agents Properties of Copper Copper Proteins Inorganic Drugs in Deficiency and Disease Zinc and Its Role in Biology and Nutrition Methods Involving Metal Ions and Complexes in Clinical Chemistry Calcium and Its Role in Biology Circulation of Metals in the Environment Antibiotics and Their Complexes Concepts on Metal Ion Toxicity Applications of Nuclear Magnetic Resonance to Paramagnetic Species ENDOR, EPR, and Electron Spin Echo for Probing Coordination Spheres Nickel and Its Role in Biology

xx

Volume 24: Volume 25: Volume 26: Volume 27: Volume 28: Volume 29: Volume 30: Volume 31: Volume 32: Volume 33: Volume 34: Volume 35: Volume 36: Volume 37: Volume 38: Volume 39: Volume 40: Volume 41: Volume 42: Volume 43: Volume 44:

VOLUMES IN THE MIBS SERIES

Aluminum and Its Role in Biology Interrelations among Metal Ions, Enzymes, and Gene Expression Compendium on Magnesium and Its Role in Biology, Nutrition, and Physiology Electron Transfer Reactions in Metalloproteins Degradation of Environmental Pollutants by Microorganisms and Their Metalloenzymes Biological Properties of Metal Alkyl Derivatives Metalloenzymes Involving Amino Acid-Residue and Related Radicals Vanadium and Its Role for Life Interactions of Metal Ions with Nucleotides, Nucleic Acids, and Their Constituents Probing Nucleic Acids by Metal Ion Complexes of Small Molecules Mercury and Its Effects on Environment and Biology Iron Transport and Storage in Microorganisms, Plants, and Animals Interrelations between Free Radicals and Metal Ions in Life Processes Manganese and Its Role in Biological Processes Probing of Proteins by Metal Ions and Their Low-Molecular-Weight Complexes Molybdenum and Tungsten. Their Roles in Biological Processes The Lanthanides and Their Interrelations with Biosystems Metal Ions and Their Complexes in Medication Metal Complexes in Tumor Diagnosis and as Anticancer Agents Biogeochemical Cycles of Elements Biogeochemistry, Availability, and Transport of Metals in the Environment

Contents of Volumes in the Metal Ions in Life Sciences Series edited by the SIGELs Volumes 1–4 published by John Wiley & Sons, Ltd., Chichester, UK (2006–2008) <www.Wiley.com/go/mils> and from Volume 5 on by the Royal Society of Chemistry, Cambridge, UK (since 2009) <www.rsc.org/shop/books/series/85.asp?seriesid=85>

Volume 1: 1. 2.

3.

4.

5.

6.

Neurodegenerative Diseases and Metal Ions

The Role of Metal Ions in Neurology. An Introduction Dorothea Strozyk and Ashley I. Bush Protein Folding, Misfolding, and Disease Jennifer C. Lee, Judy E. Kim, Ekaterina V. Pletneva, Jasmin Faraone-Mennella, Harry B. Gray, and Jay R. Winkler Metal Ion Binding Properties of Proteins Related to Neurodegeneration Henryk Kozlowski, Marek Luczkowski, Daniela Valensin, and Gianni Valensin Metallic Prions: Mining the Core of Transmissible Spongiform Encephalopathies David R. Brown The Role of Metal Ions in the Amyloid Precursor Protein and in Alzheimer’s Disease Thomas A. Bayer and Gerd Multhaup The Role of Iron in the Pathogenesis of Parkinson’s Disease Manfred Gerlach, Kay L. Double, Mario E. Go¨tz, Moussa B. H. Youdim, and Peter Riederer

xxii

CONTENTS OF MILS VOLUMES

7. In Vivo Assessment of Iron in Huntington’s Disease and Other Age-Related Neurodegenerative Brain Diseases George Bartzokis, Po H. Lu, Todd A. Tishler, and Susan Perlman 8. Copper-Zinc Superoxide Dismutase and Familial Amyotrophic Lateral Sclerosis Lisa J. Whitson and P. John Hart 9. The Malfunctioning of Copper Transport in Wilson and Menkes Diseases Bibudhendra Sarkar 10. Iron and Its Role in Neurodegenerative Diseases Roberta J. Ward and Robert R. Crichton 11. The Chemical Interplay between Catecholamines and Metal Ions in Neurological Diseases Wolfgang Linert, Guy N. L. Jameson, Reginald F. Jameson, and Kurt A. Jellinger 12. Zinc Metalloneurochemistry: Physiology, Pathology, and Probes Christopher J. Chang and Stephen J. Lippard 13. The Role of Aluminum in Neurotoxic and Neurodegenerative Processes Tama´s Kiss, Krisztina Gajda-Schrantz, and Paolo F. Zatta 14. Neurotoxicity of Cadmium, Lead, and Mercury Hana R. Pohl, Henry G. Abadin, and John F. Risher 15. Neurodegerative Diseases and Metal Ions. A Concluding Overview Dorothea Strozyk and Ashley I. Bush Subject Index

Volume 2: 1.

Nickel and Its Surprising Impact in Nature

Biogeochemistry of Nickel and Its Release into the Environment Tiina M. Nieminen, Liisa Ukonmaanaho, Nicole Rausch, and William Shotyk 2. Nickel in the Environment and Its Role in the Metabolism of Plants and Cyanobacteria Hendrik Ku¨pper and Peter M. H. Kroneck 3. Nickel Ion Complexes of Amino Acids and Peptides Teresa Kowalik-Jankowska, Henryk Kozlowski, Etelka Farkas, and Imre So´va´go´ 4. Complex Formation of Nickel(II) and Related Metal Ions with Sugar Residues, Nucleobases, Phosphates, Nucleotides, and Nucleic Acids Roland K. O. Sigel and Helmut Sigel 5. Synthetic Models for the Active Sites of Nickel-Containing Enzymes Jarl Ivar van der Vlugt and Franc Meyer


6. 7. 8.

9.

10. 11. 12.

13. 14.

15.

16. 17.

Urease: Recent Insights in the Role of Nickel Stefano Ciurli Nickel Iron Hydrogenases Wolfgang Lubitz, Maurice van Gastel, and Wolfgang Ga¨rtner Methyl-Coenzyme M Reductase and Its Nickel Corphin Coenzyme F430 in Methanogenic Archaea Bernhard Jaun and Rudolf K. Thauer Acetyl-Coenzyme A Synthases and Nickel-Containing Carbon Monoxide Dehydrogenases Paul A. Lindahl and David E. Graham Nickel Superoxide Dismutase Peter A. Bryngelson and Michael J. Maroney Biochemistry of the Nickel-Dependent Glyoxylase I Enzymes Nicole Sukdeo, Elisabeth Daub, and John F. Honek Nickel in Acireductone Dioxygenase Thomas C. Pochapsky, Tingting Ju, Marina Dang, Rachel Beaulieu, Gina Pagani, and Bo OuYang The Nickel-Regulated Peptidyl-Prolyl cis/trans Isomerase SlyD Frank Erdmann and Gunter Fischer Chaperones of Nickel Metabolism Soledad Quiroz, Jong K. Kim, Scott B. Mulrooney, and Robert P. Hausinger The Role of Nickel in Environmental Adaptation of the Gastric Pathogen Helicobacter pylori Florian D. Ernst, Arnoud H. M. van Vliet, Manfred Kist, Johannes G. Kusters, and Stefan Bereswill Nickel-Dependent Gene Expression Konstantin Salnikow and Kazimierz S. Kasprzak Nickel Toxicity and Carcinogenesis Kazimierz S. Kasprzak and Konstantin Salnikow Subject Index

Volume 3: 1.

xxiii

The Ubiquitous Roles of Cytochrome P450 Proteins

Diversities and Similarities of P450 Systems: An Introduction Mary A. Schuler and Stephen G. Sligar 2. Structural and Functional Mimics of Cytochromes P450 Wolf-D. Woggon 3. Structures of P450 Proteins and Their Molecular Phylogeny Thomas L. Poulos and Yergalem T. Meharenna 4. Aquatic P450 Species Mark J. Snyder


xxiv

5. 6. 7.

8. 9.

10.

11.

12. 13. 14.

15.

16.

17.

The Electrochemistry of Cytochrome P450 Alan M. Bond, Barry D. Fleming, and Lisandra L. Martin P450 Electron Transfer Reactions Andrew K. Udit, Stephen M. Contakes, and Harry B. Gray Leakage in Cytochrome P450 Reactions in Relation to Protein Structural Properties Christiane Jung Cytochromes P450. Structural Basis for Binding and Catalysis Konstanze von Ko¨nig and Ilme Schlichting Beyond Heme-Thiolate Interactions: Roles of the Secondary Coordination Sphere in P450 Systems Yi Lu and Thomas D. Pfister Interactions of Cytochrome P450 with Nitric Oxide and Related Ligands Andrew W. Munro, Kirsty J. McLean, and Hazel M. Girvan Cytochrome P450-Catalyzed Hydroxylations and Epoxidations Roshan Perera, Shengxi Jin, Masanori Sono, and John H. Dawson Cytochrome P450 and Steroid Hormone Biosynthesis Rita Bernhardt and Michael R. Waterman Carbon-Carbon Bond Cleavage by P450 Systems James J. De Voss and Max J. Cryle Design and Engineering of Cytochrome P450 Systems Stephen G. Bell, Nicola Hoskins, Christopher J. C. Whitehouse, and Luet L. Wong Chemical Defense and Exploitation. Biotransformation of Xenobiotics by Cytochrome P450 Enzymes Elizabeth M. J. Gillam and Dominic J. B. Hunter Drug Metabolism as Catalyzed by Human Cytochrome P450 Systems F. Peter Guengerich Cytochrome P450 Enzymes: Observations from the Clinic Peggy L. Carver Subject Index

Volume 4: 1.

Biomineralization. From Nature to Application

Crystals and Life: An Introduction Arthur Veis 2. What Genes and Genomes Tell Us about Calcium Carbonate Biomineralization Fred H. Wilt and Christopher E. Killian


3. 4.

5.

6. 7. 8. 9.

10.

11.

12. 13. 14.

15. 16.

17. 18.

The Role of Enzymes in Biomineralization Processes Ingrid M. Weiss and Fre´de´ric Marin Metal–Bacteria Interactions at Both the Planktonic Cell and Biofilm Levels Ryan C. Hunter and Terry J. Beveridge Biomineralization of Calcium Carbonate. The Interplay with Biosubstrates Amir Berman Sulfate-Containing Biominerals Fabienne Bosselmann and Matthias Epple Oxalate Biominerals Enrique J. Baran and Paula V. Monje Molecular Processes of Biosilicification in Diatoms Aubrey K. Davis and Mark Hildebrand Heavy Metals in the Jaws of Invertebrates Helga C. Lichtenegger, Henrik Birkedal, and J. Herbert Waite Ferritin. Biomineralization of Iron Elizabeth C. Theil, Xiaofeng S. Liu, and Manolis Matzapetakis Magnetism and Molecular Biology of Magnetic Iron Minerals in Bacteria Richard B. Frankel, Sabrina Schu¨bbe, and Dennis A. Bazylinski Biominerals. Recorders of the Past? Danielle Fortin, Sean R. Langley, and Susan Glasauer Dynamics of Biomineralization and Biodemineralization Lijun Wang and George H. Nancollas Mechanism of Mineralization of Collagen-Based Connective Tissues Adele L. Boskey Mammalian Enamel Formation Janet Moradian-Oldak and Michael L. Paine Mechanical Design of Biomineralized Tissues. Bone and Other Hierarchical Materials Peter Fratzl Bioinspired Growth of Mineralized Tissue Darilis Sua´rez-Gonza´lez and William L. Murphy Polymer-Controlled Biomimetic Mineralization of Novel Inorganic Materials Helmut Co¨lfen and Markus Antonietti Subject Index

xxv


xxvi

Volume 5: 1. 2. 3. 4. 5. 6. 7. 8.

9.

10. 11.

12.

13. 14.

15.

Metallothioneins and Related Chelators

Metallothioneins: Historical Development and Overview Monica Nordberg and Gunnar F. Nordberg Regulation of Metallothionein Gene Expression Kuppusamy Balamurugan and Walter Schaffner Bacterial Metallothioneins Claudia A. Blindauer Metallothioneins in Yeast and Fungi Benedikt Dolderer, Hans-Ju¨rgen Hartmann, and Ulrich Weser Metallothioneins in Plants Eva Freisinger Metallothioneins in Diptera Silvia Atrian Earthworm and Nematode Metallothioneins Stephen R. Stu¨rzenbaum Metallothioneins in Aquatic Organisms: Fish, Crustaceans, Molluscs, and Echinoderms Laura Vergani Metal Detoxification in Freshwater Animals. Roles of Metallothioneins Peter G. C. Campbell and Landis Hare Structure and Function of Vertebrate Metallothioneins Juan Hidalgo, Roger Chung, Milena Penkowa, and Milan Vasˇa´k Metallothionein-3, Zinc, and Copper in the Central Nervous System Milan Vasˇa´k and Gabriele Meloni Metallothionein Toxicology: Metal Ion Trafficking and Cellular Protection David H. Petering, Susan Krezoski, and Niloofar M. Tabatabai Metallothionein in Inorganic Carcinogenesis Michael P. Waalkes and Jie Liu Thioredoxins and Glutaredoxins. Functions and Metal Ion Interactions Christopher Horst Lillig and Carsten Berndt Metal Ion-Binding Properties of Phytochelatins and Related Ligands Aure´lie Devez, Eric Achterberg, and Martha Gledhill Subject Index

Volume 6:

Metal-Carbon Bonds in Enzymes and Cofactors (this book)


Volume 7:

1. 2.

3.

4. 5. 6.

7. 8. 9. 10.

11. 12. 13.

14.

xxvii

Organometallics in Environment and Toxicology (tentative contents)

Organometal(loid) Compounds in Environmental Cycles John S. Thayer Analysis of Organometallic Compounds in Environmental and Biological Samples Richard O. Jenkins and Chris F. Harrington Evidence for Organometallic Intermediates in Bacterial Methane Formation Involving the Nickel Coenzyme F430 Stephen W. Ragsdale and Mishtu Dey Tinorganyls. Formation, Use, Speciation, and Toxicology Tamas Gajda Alkyl-Lead Compounds and Their Environmental Toxicology Henry G. Abadin and Hana R. Pohl Organoarsenicals. Distribution and Transformation in the Environment Kenneth J. Reimer Organoarsenicals. Toxicity and Carcinogenicity Elke Dopp, Andrew D. Kligerman, and Roland A. Diaz-Bone Alkyl Derivatives of Antimony in the Environment Montserrat Filella Alkyl Derivatives of Bismuth in Environmental and Biological Media Montserrat Filella Formation, Occurrence, and Significance of Organoselenium and Organotellurium Compounds in the Environment Dirk Wallschla¨ger and Jo¨rg Feldmann Organomercurials. Their Formation and Role in the Environment Holger Hintelmann Toxicology of Alkyl-Mercury Compounds Michael Aschner Environmental Bioindication and Bioremediation of Organometal(loid)s John S. Thayer Alkylated Metal(loid) Species in Humans Alfred V. Hirner and Albert V. Rettenmeier Subject Index

Comments and suggestions with regard to contents, topics, and the like for future volumes of the series are welcome.

Met. Ions Life Sci. 2009, 6, 1–51

1 Organometallic Chemistry of B12 Coenzymes Bernhard Kraütler Institute of Organic Chemistry and Centre of Molecular Biosciences, University of Innsbruck, A-6020 Innsbruck, Austria

ABSTRACT 2 1. INTRODUCTION 2 2. STRUCTURE OF B12 DERIVATIVES IN THE CRYSTAL AND IN SOLUTION 5 2.1. ‘‘Incomplete’’ Corrinoids 5 2.2. ‘‘Complete’’ Corrinoids 7 2.2.1. ‘‘Complete’’ Corrinoids with an ‘‘Inorganic’’ b-Ligand 7 2.2.2. ‘‘Complete’’ Corrinoids with an ‘‘Organic’’ b-Ligand 10 2.2.3. Spectroscopic Studies of the Solution Structure of B12 Derivatives 13 2.3. The ‘‘Base-On/Base-Off’’ Constitutional Switch of ‘‘Complete’’ Corrinoids 16 18 3. REDOX CHEMISTRY OF B12 DERIVATIVES 3.1. Thermodynamics of Redox Processes 20 3.2. Kinetics of the Redox Processes 22 4. REACTIVITY OF B12 DERIVATIVES IN ORGANOMETALLIC REACTIONS 24 4.1. Formation of the (Co–C) Bond in Organocorrinoids 25 4.2. Cleavage of the (Co–C) Bond in Organocorrinoids 31 5. ORGANOMETALLIC B12 DERIVATIVES AS COFACTORS AND INTERMEDIATES IN ENZYMES 34 35 5.1. Methylcorrinoids in B12-Dependent Methyltransferases 5.2. Adenosylcorrinoids in Enzymes Dependent on Coenzyme B12 38 Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00001

2

KRAÜTLER

6. CONCLUDING REMARKS AND FUTURE DIRECTIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

40 41 41 42

ABSTRACT: When coenzyme B12 was identified as organometallic derivative of vitamin B12, metal-carbon bonds were revealed to be relevant in life processes. Vitamin B12, the ‘‘antipernicious anaemia factor’’ required for human health, was isolated earlier as a crystallizable cyano-Co(III)-complex. B12 cofactors and other cobalt corrinoids play important roles not only in humans, but in the metabolism of archaea and other microorganisms, in particular. Indeed, the microorganisms are the only natural sources of the B12 derivatives. For other B12-requiring organisms the corrinoids are thus ‘‘vitamins’’. However, vitamin B12 also needs to be converted into organometallic B12-forms, which are the typical coenzymes in metabolically important enzymes. One of these, methionine synthase, catalyzes the transfer of a methyl group and its corrinoid cofactor is methylcobalamin. Another one, methylmalonyl-CoA mutase uses a reversible radical process, and coenzyme B12 (adenosylcobalamin) as its cofactor, to transform methylmalonyl-CoA into succinyl-CoA. In such enzymes, the bound B12 derivatives engage (or are formed) in exceptional organometallic enzymatic reactions, which depend upon the organometallic reactivity of the B12 cofactors. Clearly, organometallic B12 derivatives hold an important position in life and have thus attracted particular interest from the medical sciences, biology, and chemistry. This chapter outlines the unique structures of B12 derivatives and recapitulates their redox properties and their organometallic chemistry, relevant in the context of the metabolic transformation of B12 derivatives into the relevant coenzyme forms and for their use in B12-dependent enzymes. KEYWORDS: cobalt-carbon bond cobalt complex coenzyme B12 electrochemistry homolysis methyl group transfer organometallic bond radical reaction vitamin B12

1. INTRODUCTION The importance of a metal-carbon bond in enzymatic processes was first revealed in the early 1960s, when coenzyme B12 was identified as organometallic derivative of vitamin B12 [1]. Vitamin B12 was discovered and isolated 60 years ago as a crystallizable, red complex [2,3], and was revealed to be a cobalt complex of the remarkable corrin ligand, a unique member of the natural tetrapyrroles [4]. Organometallic B12 forms are the coenzymes in a variety of metabolically important enzymes. In humans, methionine synthase and methylmalonyl-CoA mutase use methylcobalamin and coenzyme B12, respectively, as their B12 cofactors [5–12]. B12 coenzymes are now known to be required in the metabolism of a broad range of organisms. However, only microorganisms have the capacity


ORGANOMETALLIC CHEMISTRY OF B12 COENZYMES

3

to biosynthesize B12 and other natural corrinoids [13,14]. For other B12dependent organisms, such as humans, B12 derivatives are thus vitamins [15]. Their functioning metabolism depends on the uptake and binding of B12 derivatives [16], on their metabolic transformation into the relevant B12 cofactors [17], the controlled transport of these [16] and the catalysis by B12-dependent enzymes [18–23]. Perhaps B12 coenzymes are Nature’s physiologically most relevant organometallic cofactors [9–12]. Organometallic B12 derivatives engage in protein-activated reactions in unique B12-dependent enzymes. In these, B12 cofactors (co)catalyze exceptional enzymatic reactions [18–23] that directly depend upon the reactivity of the cobalt-carbon bond [11,12,22]. Since the surprising identification of coenzyme B12 as an organometallic compound, corrinoid cofactors have thus played central roles in the discovery and better understanding of biological organometallic processes [24–28]. Vitamin B12 was identified as the ‘‘antipernicious anemia factor’’ over 60 years ago. It was isolated as the red cyanide-containing cobalt complex cyanocob(III)alamin (1, CNCbl), which crystallized readily and was revealed to be a relatively inert Co(III) complex [2,3]. It is the most important commercially available form of the naturally occurring B12 derivatives, but it appears to have no physiological function itself [15]. The physiologically relevant vitamin B12 derivatives are the highly light-sensitive and chemically more labile organometallic coenzymes, coenzyme B12 (2, 5 0 -deoxy-5 0 -adenosylcobalamin, AdoCbl) and methylcobalamin (3, MeCbl), as well as the ‘‘inorganic’’ and easily reducible B12 derivatives aquacob(III)alamin (41, H2OCbl1) and hydroxocob(III)alamin (5, HOCbl) [6,29,30] (Figure 1). During the last six decades, remarkable scientific advances towards the solution of some of the major ‘‘B12-mysteries’’ have been achieved. Five European Symposia on ‘‘Vitamin B12 and B12-Proteins’’ were dedicated to this subject, the first two of which took place in Hamburg (1956 and 1961, Germany), followed by one in Zu¨rich (1979, Switzerland) [7], in Innsbruck (1996, Austria) [9], and in Marburg (2000, Germany). Among the top achievements in the B12 field the elucidation of the structure of vitamin B12 [4,31] and of coenzyme B12 [1,32] are to be highlighted, the synthetic conquest of the vitamin B12 structure [33–35], the elucidation of the biosynthetic pathways to B12 [13,14], as well as crystal structures [36–44] and a solution structure [45] of a variety of B12-binding proteins and B12-dependent enzymes. Several concise books on B12 have been written, with earlier ones by Pratt [5] and by Friedrich [6]. More recent ones on ‘‘B12’’ [7], ‘‘Vitamin B12 and B12-Proteins’’ [9] and on ‘‘Chemistry and Biochemistry of B12’’ [10] and extensive reviews [11,12] describe the more recent findings on the chemistry of B12 and on the biological roles of the B12 derivatives.


KRAÜTLER

4 H2NOC

H2NOC CONH2 CH3

H2NOC

CONH2 CH3

H2NOC

CH3

CH3 CONH2

CONH2 H3C

N

H3C

R

Co+ N N

H

H 3C

N

H2NOC

N

Co+ N

H3 C H CH3 CH3

H3C

N

R

N

H2NOC

CH3

H 3C

CH3

CH3 CONH2

CONH2 O

HN

CH3

N

O

HN

Nu

H 3C

H3C HO

H O

N

CH3 O

O

N

HO O

O

O

N

N

NH2

P -O

O

N

Nu =

H

P -O

CH3

O

OH

OH R

R

Co+

Co+

N N

N

CH3 N

NH2 N N

-O

DMB

-O

Nu

Figure 1. General structural formula. Left: cobalamins (Cbls ¼ DMB-cobamides, Ado ¼ adenosyl). Vitamin B12 (1, CNCbl, R ¼ CN), coenzyme B12 (2, R ¼ 5 0 -deoxy5 0 -ado), methylcobalamin (3, MeCbl, R ¼ CH3), aquacobalamin (41, R ¼ H2O1), hydroxocobalamin (5, HOCbl, R ¼ HO), cob(II)alamin (6, B12r, R ¼ e), chlorocobalamin (18 , R ¼ Cl), nitroxylcobalamin (19, R ¼ NO), 2,3-dihydroxypropyl-Cbl (21, R ¼ 2,3-dihydroxy-propyl), a-adenosyl-Cbl (22, R ¼ 5 0 -deoxy-5 0 -a-Ado), adeninylpropyl-Cbl (23, R ¼ 3-adeninyl-propyl), homocoenzyme B12 (24, R ¼ 5 0 -deoxy-5 0 Ado-methyl), trifluoromethyl-Cbl (25, R ¼ CF3), difluoromethyl-Cbl (26, R ¼ CHF2), vinylcobalamin (28, R ¼ CH¼CH2), cis-chlorovinyl-Cbl (29, R ¼ CH¼CHCl), bishomocoenzyme B12 (33, R ¼ 2-[5 0 -deoxy-5 0 -Ado]-ethyl), 2 0 -deoxycoenzyme B12 (48, R ¼ 2 0 ,5 0 -dideoxy-5 0 -Ado), 2 0 ,3 0 -dideoxycoenzyme B12 (49, R ¼ 2 0 ,3 0 ,5 0 -trideoxy-5 0 -Ado). Right: Structural formulae of other naturally occurring ‘‘complete’’ corrinoids (cobamides with other nucleotide functions ‘‘Nu’’ [6,11]: Cobcyano-imidazolylcobamide (14, R ¼ CN, Nu ¼ imidazole), Cob-methyl-imidazolylcobamide (27, R ¼ CH3, Nu ¼ imidazole); pseudovitamin B12 (Cob-cyano-700 -adeninylcobamide, 16, R ¼ CN, Nu ¼ adenine), factor A (Cob-cyano-700 -[2 0 -methyl]adeninyl-cobamide, 17, R ¼ CN, Nu ¼ 2-methyl-adenine). Bottom: symbols used.



5

2. STRUCTURE OF B12 DERIVATIVES IN THE CRYSTAL AND IN SOLUTION 2.1. ‘‘Incomplete’’ Corrinoids The structures of vitamin B12 (1) and of coenzyme B12 (2) were established largely through the pioneering X-ray crystallographic studies of Hodgkin et al. [4,31,32], who discovered the composition of the corrin core of 1 and the organometallic nature of 2 (see recent reviews [46–48]). These two cobalamins belong to the ‘‘complete’’ corrinoids, in which a pseudonucleotide function is attached via an amide linkage to the corrin moiety. The resulting combination is unique, as the B12-nucleotide function may switch between the cobalt-coordinated ‘‘base-on’’ form and the de-coordinated (so called ‘‘base-off’’ form) – coenzyme B12 thus can be considered to be a ‘‘molecular switch’’ [49]. Earlier important X-ray investigations also specifically relied on some crystallizable ‘‘incomplete’’ Co(III)-corrinoids, such as a-cyano-b-aqua cobyric acid (7) (see review [46]). Cobyric acid is the natural corrinoid moiety of the vitamin B12 and it was the initial target for Eschenmoser and Woodward for the total synthesis of vitamin B12 [33–35] (as it had already been shown how 7 could be converted to vitamin B12). Since these times, crystallographic work with ‘‘incomplete’’ Co(III)-corrinoids has focused on obtaining detailed structural information. More recently analyzed structures include that of dicyano-heptamethyl-cobyrinate (8, ‘‘cobester’’ [50]) [51,52] and of its analogues [53], including 15-norcobester (9) [54,55] (as reviewed in [47]). Very recently, a new type of B12 dimer structure was found in the crystal of Cob-cyano-neocobyric acid (10) [56] (Figure 2). A similar dimer structure has not been observed in the ‘‘normal’’ corrinoids. However, in the ‘‘neo’’ corrinoid 10, the configuration at the corrin ring C was inverted, apparently reducing the steric hindrance due to a propionate side chain, and making the dimer formation possible [56]. The crystal structure of heptamethyl-cob(II)yrinate (cob(II)ester (as the perchlorate complex 11) gave the first detailed insights into the structure of a paramagnetic Co(II)-corrin [57]. It revealed a five-coordinated Co(II) center in the ‘‘incomplete’’ Co(II)-corrin 11, to which a perchlorate ligand was coordinated via a long axial cobalt-oxygen bond (2.31 A˚) [57]. The coordination of the axial ligand at the sterically less hindered ‘‘upper’’ b-face is in contrast with the preference seen in the ‘‘complete’’ cob(II)alamin (6, B12r) [58,59] (see below).


KRAÜTLER

6 H2NOC

H3CO2C

CONH2

H2NOC

CH3

H3C H3C

N

R

CO2CH3 H3CO2C

CH3

H3C

CONH2

N N

N L′

H2NOC H3C

H

CH3 13

CH3

R1

CH3

H3CO2C

R1′

H3C

CO2CH3

Co N

L′

N

CH3 CH3

X CO2CH3

H3CO2C

X

O

CH3

N L N

H3C

Co+

H

CH3

O34 N34

C72

C32

C22 N23

O73

C33

O23

C51

C31

C21

C3

C5 C4

C71

N73

C7A

C6 C7

C2 C8 C81 N84 R N1 C2A N2 C9 C1 C1A C82 C83 Co C10 C19 C181 O84 N3 C11 N4 C12B O183 L C18 C12 C14 C16 C182 C12A C17 C13 C15 C17B N183 C171 C131 O C151 C172 C132

+

N174

O134

CoIII

L NH H CH3

C133

C173 O174

R

N134

OH

C175 C176 C177

O177

Figure 2. Top left: Structural formulae of ‘‘incomplete’’ corrinoids: Coa-cyano-Cobaqua cobyric acid (7, R ¼ H2O, R1 ¼ propionamide, R01 ¼ H, L 0 ¼ CN, X ¼ OH); Coa-aqua-Cob-cyano-13-epicobyric acid (10, R ¼ CN, R01 ¼ propionamide, R1 ¼ H, L 0 ¼ H2O, X ¼ OH), Cob-5 0 -deoxy-5 0 -adenosylcobinamide (32, R ¼ 5 0 -deoxy-5 0 -Ado, R1 ¼ propionamide, R01 ¼ H, L 0 ¼ H2O, X ¼ NH–CH2–CHOH–CH3). Top right: Structural formulae of heptamethyl-cobyrinates: cobester (8, L ¼ L 0 ¼ CN, X ¼ CH3), 15-nor-cobester (9, L ¼ L 0 ¼ CN, X ¼ H); perchlorato-heptamethylcob(II)yrinate (11, L ¼ ClO4, L 0 ¼ vacant, X ¼ CH3). Bottom left: Atom numbering for cobinamide [60]. Bottom right: general symbol used for a cob(III)inamide.



7

2.2. ‘‘Complete’’ Corrinoids The ‘‘complete’’ corrinoid vitamin B12 (1, CNCbl) is called a ‘‘cobalamin’’ (or a 5 0 ,6 0 -dimethylbenzimidazolyl-cobamide ¼ DMB-cobamide), in which a cyanide ligand is bound at the ‘‘upper’’ axial coordination site (on the b-face, see e.g., [60]). In other ‘‘cobalamins’’, the cyanide group at the b-face of CNCbl is replaced by a different ligand, e.g., an organometallic group, as in coenzyme B12 (AdoCbl, 2). Purinyl-cobamides are another important class of naturally occurring ‘‘complete’’ corrinoids, in which a purine base is part of the nucleotide function (see Figure 1) [6,11]. The systematic atom numbering used in this chapter for vitamin B12 derivatives [60] builds on the convention that atom numbers of the heavy atoms of a substituent reflect the number of the points of attachment to the corrin ligand and are indexed consecutively [61] (see Figure 2); however, it deviates from that introduced by Hodgkin et al. [31,32,46] and still used by some authors [12].

2.2.1. ‘‘Complete’’ Corrinoids with an ‘‘Inorganic’’ b-Ligand Only ‘‘base-on’’ cobalamins, where the nucleotide functionality coordinates in an intramolecular mode, have been analyzed by X-ray crystallography [46–48]. The ‘‘old’’ structure of vitamin B12 [7] was re-analyzed using modern cryo-crystallography techniques [47,62], which showed the molecular geometry of the B12 moiety to agree within experimental error of Hodgkin’s original result. Neovitamin B12 (12) was also studied by crystallography and was revealed to be the cyano-13-epicobalamin, a derivative of vitamin B12 where the propionamide chain at the C13 position is in the b-configuration [63,64]. A notable difference between the two structures is an increased ‘‘nonplanarity’’ in the corrin ring of the neo-derivative (expressed as a 61 larger fold angle, 23.71 versus 17.91). The C8 epimer of vitamin B12, cyano-8-epicobalamin (13), has an even larger fold angle of the corrin core (23.81) [65] (Figure 3). The discovery of the replacement of the cobalt-coordinated 5,6-dimethylimidazole base by a protein-derived imidazole in several B12-dependent enzymes [36–38]), gave the analysis of Cob-cyano-imidazolylcobamide (14) [62] particular interest. The less bulky and more nucleophilic imidazole base of 14 caused a number of structural differences. The corrin ring fold angle of 14 decreased to 11.31 and the axial (Co–N) bond shortened (from 2.011 A˚ in 1 to 1.968 A˚ in 14). In addition, the ‘‘base tilt’’ of 12 (i.e., half the difference between the two Co–N–C angles to the coordinating base) decreased to near zero, within experimental error. In all crystal structures of 5 0 ,6 0 -dimethylbenzimidazoyl-cobamides a ‘‘tilt’’ of about 51 is Met. Ions Life Sci. 2009, 6, 1–51


-O

O

P

HN

O

HO

O

O

H3C

H

N

N

R2

OH

O

CH3

13

8

CH3

CH3

R2′

CH3

CH3

R1′ R1

CH3

CONH2

H

R′

-O

O P O

CH3

OH

O

Nu

CH3

Co+ N N

N CN N

HO

O

O

H3C

HN

H2NOC

H

H3C

H3C

H2NOC

H2NOC

CONH2

CH3

CH3

CH3

CONH2

H

-O

O P

O

O

O

H3C

HN

H2NOC

H

H3C

H3C

H2NOC

H3C

CONH2

H2NOC

OH

N

O

OH

CH3

Co+ N L

N R N

CH3

Nu

CONH2

CH3

CH3

CH3

CONH2 CONH2

Figure 3. Left: Structural formulae of neovitamin B12 (12, cyano-13-epicobalamin, R ¼ CN, R01 ¼ R2 ¼ H, R1 ¼ R02 ¼ propionamide), cyano-8-epi-cobalamin (13, R ¼ CN R1 ¼ R02 ¼ H, R01 ¼ R2 ¼ propionamide), neocoenzyme B12 (36, R ¼ 5 0 -deoxy-5 0 -adenosyl, R01 ¼ R2 ¼ H, R1 ¼ R02 ¼ propionamide). Center: Structural formulae of pseudovitamin B12 (16, R 0 ¼ CH3; Nu ¼ adenine), norpseudovitamin B12 (15, R 0 ¼ H; Nu ¼ adenine), norvitamin B12 (37, R 0 ¼ H; Nu ¼ DMB). Right: Structural formulae of base-off cobamides: pseudocoenzyme B12 (34, R ¼ 5 0 -deoxy-5 0 -adenosyl; L ¼ H2O; Nu ¼ adenine), adenosyl-factor A (35, R ¼ 5 0 -deoxy-5 0 -adenosyl, L ¼ H2O, Nu ¼ 2-methyl-adenine), cob(I)alamin– (B 12s, 39 , R ¼ L ¼ absent, Nu ¼ DMB).

H

H3C

H2NOC

H3C

Co+ N N

N R N

H2NOC

H3C

CH3

H2NOC

8 KRAÜTLER


9

found [47,62], which appears to be an inherent property of the cobaltcoordinated DMB. Norpseudovitamin B12 (Cob-cyano-700 -adeninyl-176-norcobamide) (15) represented the first natural example of a ‘‘complete’’ B12 derivative that lacked one of the methyl groups (of C176) of the cobamide moiety [66]. Xray crystal structures were determined for 15 and for the analogues, pseudovitamin B12 (16) and factor A (Cob-cyano-700 -[2-methyl] adeninylcobamide) (17) [66]. These first accurate crystal structures of complete corrinoids with an adeninyl pseudo-nucleotide confirmed the expected coordination properties around Co and corroborated the virtual conformational identity of the nucleotide moieties of 15 and its two homologues. For 16, an axial (Co–N) bond of 2.026 A˚ and a fold angle of 16.91 were determined [66], for 15 and 17, the axial (Co–N) bonds were 2.035 A˚ and 2.021 A˚ respectively, and both had a fold angle of 19.61. The observed structural changes from the replacement of DMB by adenine or 2-methyladenine in cyano-Co(III) cobamides are hardly significant. In line with this structural point of view, purinyl-cobamides, such as 16 and 17, have been shown recently to belong to a broadly occurring and naturally biosynthesized type of ‘‘complete’’ corrinoids [67]. For the crystal structure of aquacobalamin perchlorate (41-ClO 4 ) [68] the shortest known axial (Coa–N) bond of a vitamin B12 derivative was observed (1.925 A˚). Together with the large upwards folding angle of 18.71, the conclusion stated was, that steric repulsion between the DMB base and corrin core led to a flexing of the corrin ring [47,68]. The short axial Coa–N bond for the 41-ion was consistent with the weak donor ability of the (trans-axial) Cob-aqua ligand. Crystal structures of various other ‘‘inorganic’’ B12 derivatives have been solved and previously reviewed elsewhere [46–48]. For several other ‘‘inorganic’’ B12 derivatives crystal structures were obtained recently, including chlorocobalamin (18) [69] and NO–Cbl (19), which is best described a nitroxyl-cob(III)alamin [70]. Crystal structures of a cobalamin derivative, where the cyanide ligand of vitamin B12 acts as bridging ligand between the rhenium carbonyl compounds (i.e., with a central Co–CN–Re feature) have been analyzed [71]. The attachment of diagnostically active ligands to a cobalamin has been explored as a way of using vitamin B12 derivatives (as a ‘‘Trojan Horse’’) for introducing fluorescent [72] and radio-labeled compounds [73], or other bioactive molecules [74], into cells and living animals. Information on cob(II)alamin (6, B12r) was of particular interest [58], as (in a formal sense) it is the product of (Co–C) bond homolysis of coenzyme B12 (2), and occurs during the catalytic cycle of coenzyme B12-dependent enzymes. The crystal structure of cob(II)alamin showed that the corrin moiety of 6 and 2 were very similar [58]. The fold angle of the corrin ring in 6 is 16.31 compared to 13.31 in 2. The axial cobalt-nitrogen bond is even Met. Ions Life Sci. 2009, 6, 1–51

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slightly shorter at the five coordinate Co(II) ion of cob(II)alamin (2.13 A˚) than in the six-coordinated case in coenzyme B12 (2.24 A˚). However, the distance between the corrin ring and the coordinated DMB base is almost the same, due to a ‘‘downward’’ displacement of the cobalt atom from the plane of the corrin ligand in 6. It was expected that the reduced Co(II) ion would have a longer bond than the Co(III) species. In view of this result, in 2 and related organocobalamins, the ‘‘structural trans effect’’ of the organic ligand appears to increase the axial Co(III)–N bond which compensates for the larger covalent radius of Co(II) compared to Co(III). From these observations the conclusion was made that the interactions (apoenzyme/ coenzyme) at the corrin moiety of the coenzyme appear to be inadequate to provide the major means for a protein-induced activation of the bound coenzyme toward homolysis of its (Co–C) bond. Instead, the organometallic bond may be labilized by way of apoenzyme (and substrate) induced separation of the homolysis fragments, made possible by strong binding of the separated components to the protein [58]. Cob(II)alamin (6) has also been recently studied using neutron Laue diffraction studies, which came to the same conclusions regarding its structure [59]. In single crystals, cob(II)alamin (6) can be loaded with molecular oxygen at low temperature, to give a well ordered crystalline oxygenated complex (investigated earlier by ESR spectroscopy [75]) and best described as superoxocob(III)alamin (20), according to the X-ray analysis of the crystal structure [76]. In contrast, in (aqueous) solution, B12r (6) is readily oxidized by air to aqua-cob(III)alamin (4).

2.2.2. ‘‘Complete’’ Corrinoids with an ‘‘Organic’’ b-Ligand The original crystal structure of coenzyme B12 (2) primarily helped to reveal the organometallic nature of 2 [1,32,77]. The organometallic adenosyl moiety was observed to be bound in an anti conformation and the adenine ring was found to be above ring C of the corrin ligand. However, this and more extensive recent studies by X-ray and neutron crystallography [78,79] also showed both axial (Co–C) (2.030 A˚) and (Co–N) (2.237 A˚) bonds to be relatively long [46,47]. In addition, the organometallic group in 2 exhibited a strikingly large Co–C5 0 –C4 0 bond angle of 125.41 [46–48,77]. To investigate if the large Co–C–C bond angle of AdoCbl (2) is typical for organocobalamins, the crystal structures of the 2,3-dihydroxypropylcobalamins (the diastereomeric R- and S-isomers 21R and 21S) were examined earlier [80]. The (Co–C) distances (2.00 and 2.08 A˚ for 21R and 21S, respectively) were similar to AdoCbl (2.03 A˚) but the bond angles were smaller (119.61 for 21R and 113.61 for 21S). The value for 21S should be Met. Ions Life Sci. 2009, 6, 1–51


11

considered the ‘‘normal’’ angle, with little interactions between the corrin ring and b-substituent. In a-adenosylcobalamin (22), a stereoisomer of AdoCbl (2), the organometallic adenine base is attached at the ribose moiety in an a-configuration. The crystal structure of 22 showed the lengths of the axial (Co–C) (2.02 A˚) and (Co–N) (2.24 A˚) bonds to be similar to 2 but the corrin ring was flatter (fold angle ¼ 11.71 versus 13.31 in 2) [81]. As in 2, the adenosyl ligand was placed over the south-east quadrant (ring C), but the position of the adenine moiety relative to the ribose unit of the organometallic ligand was disordered due to different conformations of the adenine heterocycle. Adeninylalkylcobalamins, where a methylene chain connects the adenine with the cobalt center [82], inhibit various AdoCbl-dependent enzymes depending upon the length of the alkyl chain [83]. 3-Adeninylpropylcobalamin (23) has been studied in solution as well as in the crystal [84]. The structure of the corrin ring and the lower nucleotide loop closely resembled that of 2. However, the adenine group of 23 is oriented almost parallel to the corrin plane and is positioned over ring D of the corrin ligand, i.e., about 120 1 clockwise from its position in coenzyme B12. The homologue of coenzyme B12 ‘‘homocoenzyme B12’’ (24, Cob-(5 0 deoxy-5 0 -adenosylmethyl)-cob(III)alamin) has been recently prepared, as it has been suggested to function as a covalent structural mimic of the hypothetical enzyme bound ‘‘activated’’ state of the B12 cofactor [85]. In the crystal structure of 24 the cobalt center was observed to be at a distance of 2.99 A˚ from C5 0 of the homoadenosine moiety and the latter to be in the unusual syn conformation. In 24 the crucial distance from the corrin-bound cobalt center to the C5 0 of the homoadenosine moiety is, thus, roughly the same as found in one of the two ‘‘activated’’ forms of coenzyme B12 in the crystal structure of glutamate [86]. Indeed, ‘‘homocoenzyme B12’’ (24) is bound intact to glutamate mutase and does not function as cofactor [87]. In contrast, dioldehydratase and ethanolamine lyase, when reconstituted with 22, still show weak activity [88]. The crystal structure of methylcobalamin (MeCbl, 3), the simplest organometallic B12-derivative, was analyzed in 1985 [89]. Interestingly, the solution structure of MeCbl, as derived by NMR spectroscopy [90], deviated from the earlier crystal with respect to the conformation of the nucleotide loop. The structures of crystals of MeCbl (from various solvent compositions) were thus re-investigated more recently, to provide a more accurate picture [91]. The structures confirmed the folding of the corrin core of 3 to be similar to that of coenzyme B12 (AdoCbl, 2) (fold angle in 3 ¼ 14.71 [91]). This proved that the bulkiness of the 5 0 -deoxyadenosyl ligand in 2 was not a main contributor to the conformation of the corrin ligand of AdoCbl. The lengths of the axial (Co–C) (1.979 A˚) and (Co–N) (2.162 A˚) bonds are slightly shorter in MeCbl when compared to AdoCbl. The shorter axial bond Met. Ions Life Sci. 2009, 6, 1–51

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to the DMB base is consistent with the stronger nucleotide coordination in 3. The crystal structures of the fluorinated analogues of MeCbl, of CF3cobalamin (25) [92] and CF2H-cobalamin (26) were analyzed likewise [93]. Cob-methyl-imidazolylcobamide (27) was prepared as a model for methylcorrinoid cofactors in a ‘‘base-off/His-on’’ form and its crystal structure was analyzed [94]. The substitution by a less bulky and more nucleophilic imidazole base had the expected structural effects. The axial (Co–C) (1.97 A˚) and (Co–N) (2.09 A˚) bonds are shorter in 27 than in methylcobalamin and the fold angle of the corrin ligand was reduced by over 21 to 12.51. The structures of vinylcobalamin (28) [95] and cis-chlorovinylcobalamin (29) [96] were analyzed in the crystal as the first examples of organo-cobalamins with sp2-hybridized carbon ligands. The cobalamin 29 is considered a model for a putative intermediate in the reductive degradation of chlorinated ethylenes [97]. As expected for a vinyl ligand, the Co–C bond length (1.912 A˚ for 28 and 1.952 A˚ for 29) is shorter than in adenosylcobalamin (2). Steric repulsion presumably causes the significantly longer (Co–C) bond in 29.The axial (Co–N) bond lengths of 2.166 A˚ for 28 and 2.144 A˚ for 29 are also slightly shorter than in 2 and provide a good example of the ‘‘inverse’’ trans effect. Crystallography has also contributed to the elucidation of the mutual arrangement of the two closely placed corrinoid moieties in the tetramethylene-bridged organometallic B12-dimer tetramethylene-1,4-di-Cobcobalamin (30, see Figure 9 in Section 4.1) [98]. Likewise, the mode of bonding in the sterically strained conjugate (31) of MeCbl (3) and thymidine was revealed by the crystal structure of the remarkable sodium complex (31) of the ‘‘base-on’’ form of this organometallic ‘‘complete’’ corrinoid [49]. The ‘‘thermodynamic’’ and ‘‘structural’’ trans effects of B12 derivatives are the effect of one cobalt-coordinated axial ligand on chemical equilibria and coordination properties of an axial ligand trans to the first one [99]. An increasing s-donor power of the Cob ligand X was found to correlate with the size of the thermodynamic trans effect in B12 derivatives. The length of the axial (Coa–N) bond to the DMB base in cobalamins generally increases with the s-donor property of the Cob ligand [46–48]. In the same sequence, the s-ligand influences the base-on/base-off equilibria. A linear correlation thus exists between free enthalpy of the base-on/base-off equilibria in aqueous solution and the length of the (Co–N) bond [64]. However in B12 derivatives, both axial bonds lengthen simultaneously with increasing sdonor character of the axial ligands [46–48], a result of an ‘‘inverse’’ trans effect [100]. The saturated and direct trans junction between two of its four fivemembered rings is the main cause of the non-planar nature of the corrin core in B12 derivatives. The characteristic ‘‘ligand folding’’ is a main factor to the variability in the conformation of the corrin ligand [101]. The fold has Met. Ions Life Sci. 2009, 6, 1–51


13

always been found as ‘‘upwards’’ (towards the b-face), about the C10–Co axis, and the ‘‘fold angle’’ is defined as the angle between the best planes through N1–C4–C5–C6–N2–C9–C10 and C10–C11–N3–C14–C15–C16–N4 [46]. Fold angles are mostly smaller in ‘‘incomplete’’ corrinoids with a typical observed value of 7.51 in dicyano-heptamethyl-cob(III)yrinate [52], when compared to ‘‘complete’’ corrinoids, where a value of 17.91 has been found in CNCbl (1, [47]). However, in epi-corrinoids the fold angle was larger both in the ‘‘complete’’ corrinoid cyano-8-epicobalamin (13) (23.81) [65], and in the ‘‘incomplete’’ cyano-aqua-neo-cobyric acid (10) (22.51) [56]. In cobalamins, the bulky DMB base was thus suggested to be a relevant contributor to the upwards folding of corrins [46,47]. This possible effect of the intramolecular coordination of the DMB base on the folding of the corrin in Cob(III)alamins has been examined in several model situations [47,62]. Both ‘‘inorganic’’ and ‘‘organometallic’’ cob(III)alamins have been compared and the conclusion is that longer (Coa–N) bonds correlate with smaller ‘‘fold’’ angles (and vice versa) [47]; for example, aquacobalamin ˚ perchlorate (41-ClO 4 , Coa–N ¼ 1.925 A, fold angle ¼ 18.71) and coenzyme ˚ B12 (2, Coa–N ¼ 2.237 A, fold angle ¼ 13.31). In contrast, the folding of the corrin ligand in Cob-cyano-imidazolylcobamide (14) (11.31) is less than half of that of vitamin B12 (1) regardless of the shorter (Coa–N) bond (1.97 A˚ versus 2.01 A˚) [62]. Accordingly, ‘‘folding’’ is more apparent in cob(III)alamins with short (Coa–N) bonds (near 2.0 A˚ or less), to which the known ‘‘inorganic’’ B12 derivatives belong to. In organyl-cobalamins (such as methylcobalamin and coenzyme B12) the length of the (Coa–N) bond is close to or greater than 2.2 A˚, so there is less steric interaction of the nucleotide base with corrin ligand.

2.2.3. Spectroscopic Studies of the Solution Structure of B12 Derivatives Nuclear magnetic resonance (NMR) spectroscopy has had a strong influence in the development of B12 chemistry. A major feature of NMR spectroscopy is that the structures of non-crystalline B12 derivatives can be characterized in solution. The early NMR spectroscopic studies thus established the nature of many non-crystalline B12 derivatives, mostly in their Cob-cyano forms, using one-dimensional analyses [102]. These studies were based on the 1H and 13C chemical shift values from spectra of several already well-characterized B12 derivatives and used to identify and describe the structure of synthetic and natural analogues of vitamin B12 [66,103]. Along these lines, the natural corrinoids from a range of bacteria were first characterized by NMR [104,105]. Met. Ions Life Sci. 2009, 6, 1–51

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Earlier assignment problems regarding B12 derivatives in aqueous or nonaqueous solutions have now been eliminated by the use of heteronuclear NMR spectroscopy [103,106]. Following on from the pioneering studies of coenzyme B12 (2) [107,108] and the non-crystalline B12 derivative Cob-5 0 deoxy-5 0 -adenosylcobinamide (32) [109], NMR-studies in solution have begun to complement (and in certain aspects rival) X-ray analytical studies of B12 derivatives in the solid state. By applying a selection of, now wellestablished, homo- and heteronuclear 2D experiments, the assignment of signals in 1H, 13C, and 15N spectra provide a reliable basis for detailed structure and dynamic information of B12 derivatives. Techniques for suppression of the solvent (water) signal allow the recording of spectra from an aqueous solution with little or no loss of information [106]. Characteristic chemical shift values from 1H, 13C, 15N, and 31P spectra provide important information on the constitution and conformation of ‘‘complete’’ B12 derivatives [103,106]. The coordination of the DMB base, in ‘‘base-on’’ compounds, induces a high-field shift of the 1H NMR signal of HC10, due mainly to an increase in the electron density of the corrin ligand by the axial coordination of the base. This characteristic has been used to determine the temperature-dependent ‘‘base-on/base-off’’ equilibria (in aqueous solutions) of organometallic B12 derivatives (e.g., methylcobalamin 3). In the 1H NMR spectrum of, e.g., 3, the anisotropic shielding effect of the coordinated DMB base also induces high-field shift of protons located nearby, such as of the methyl group H3C1A and methylene groups H2C81 and H2C82 [90]. Shielding by the cobalt-corrin in the axial direction leads to high-field shifts of the DMB protons closest to the cobalt-corrin, HC2N and HC4N. Likewise protons of organometallic ligands are characteristically up-field as seen in the 1H NMR spectra of homocoenzyme B12 (24) and bishomocoenzyme B12 (33) [85]. Significant conformational differences between the solution and crystal structure were revealed in some cases, such as in the studies of AdoCbl (2) [107] and MeCbl (3) [90]. One of the main examples of this is the natural ‘‘complete’’ but ‘‘base-off’’ protonated form of coenzyme B12 (2-H1) [108]. More recently, the solution structures of the organometallic derivatives pseudocoenzyme B12 (34), adenosyl-factor A (35) [110], and neocoenzyme B12 (36) [111] could be analyzed in great detail (see Figure 3). The structures and ‘‘base-on/base-off’’ equilibria of a range of ‘‘complete’’ cobamides have also been studied in aqueous solution [103,106]. From NOE measurements a reliable assignment (‘‘upper’’/Cob or ‘‘lower’’/ Coa) of the cobalt-bound methyl group in non-crystalline methyl-cob(III)yrinates was achieved [112]. Also from NOE data and three bond coupling constants, detailed and important information on the conformational properties of the nucleotide moiety, the organometallic group, and of other



15

peripheral side chains was extracted [103,106]. Such studies resulted in the detection of significant conformational dynamics of the organometallic 5 0 deoxy-5 0 -adenosyl moiety in the pioneering study of coenzyme B12 (2) [107,108]. In a related context extensive conformational dynamics of the organometallic adenosyl ligand and the unusual syn orientation of the adenine heterocycle were observed in a series of coenzyme B12 analogues, such as homo- and bishomocoenzyme B12 (24 and 33) [85], pseudocoenzyme B12 (34) [110], neocoenzyme B12 (36) [111], and other adenosyl-cobamides [106]. In the solution structures of adeninyl-alkyl-cobamides, significant conformational flexibility of the organometallic ligand was also discovered [82,84]. NMR spectroscopy has proven to be a versatile method in the detection of intra- and inter-molecular H bonding. The water ligand of H2OCbl-perchlorate (41-ClO 4 ), which from the crystal structure forms an H bond to an acetamide side chain, was shown by NMR to still form a similar H bond in aqueous solution [68]. Pseudo-intramolecular H bonding of a specific ‘‘external’’ water molecule to the nucleotide portion of methylcobalamin (3) was characterized by NMR spectroscopy [90], which is accompanied by a remarkable adjustment in the conformation of the nucleotide moiety [106]. In this way first insights into the hydration behavior of B12 derivatives in aqueous solution were gained. Further exploratory studies, in which B12 derivatives were investigated in greater detail in their solvent environment, complement other recent results obtained from studies on the structure of the water networks in crystals of B12 derivatives [68,79]. The aqueous solution environments of 3 and the protonated ‘‘base-off’’ form (3-H1) of 3 have been investigated in such a way, by measuring NOEs between the solvent. Initial results also support the presence of a water molecule as the Coa-axial ligand, thus providing the first experimental evidence for a hexacoordinated cobalt center in the (solution) structure of an organometallic cobyrinic acid derivative [113]. Electron spin resonance spectroscopy has likewise provided important information on the coordination environment of Co(II)-corrins, where DMB coordination in cob(II)alamin (B12r, 6) can be detected well by ESR [114]. This technique actually also gave the first hints that Co(II) coordination of the endogenous DMB base may not be typical of enzyme-bound cob(II)alamin (6); instead, in some B12-dependent organisms histidine binding was indicated by ESR spectroscopy [115] as was found by crystallography in various B12-dependent enzymes [36,37,116]. Modern, 2D-ESR techniques have also allowed the detection of external ligands to ‘‘incomplete’’ Co(II)-corrins in (frozen) solution, such as Co(II)cobester (11) [117], and in the protonated, base-off-form of cob(II)alamin (6-H1) [118]. The application of modern methods of absorbance spectroscopy, combined with theory-based interpretation of the data has also opened a new look at


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important open questions on the coordination properties of (enzyme-bound) corrinoids (see e.g., [119,120] and Chapter 12 of this book).

2.3. The ‘‘Base-On/Base-Off’’ Constitutional Switch of ‘‘Complete’’ Corrinoids The typical functional B12 cofactors are ‘‘complete’’ corrinoids, and are unique conjugates of natural cobyrinates with unusual a-nucleotide function [6,11,12]. The constitution of the ‘‘nucleotide base’’ can vary in a remarkable way in the natural ‘‘complete’’ corrinoids. Not only benzimidazoles (such as the 5,6-dimethylbenzimidazole of the cobalamins) are among the known nucleotide functionalities, but also purines (such as adenine and 2-methyladenine in pseudovitamin B12 (16) and factor A (17), respectively) are found abundant [121]. This variety in the structure of the a(pseudo)-nucleotide unit appears to be largely a consequence of the particular biosynthetic availability in the various microorganisms [67]. The known purine bases of ‘‘complete’’ corrinoids are mostly adenine derivatives or related heterocycles [105] as also found in RNA [122]. While the DMB base is directly accessible from degradation of riboflavine in aerobes [121,123], anaerobes have evolved an unrelated and still incompletely elucidated, complex biosynthetic path to DMB and to a variety of related benzimidazoles [121]. In cobalamins, the DMB nucleotide function is structured in such a way that relatively strong intramolecular cobalt coordination is possible, and occurs with little build-up of strain [124]. However, most cobalamins and other ‘‘complete’’ corrinoids are still observed in two forms: they either have their nucleotide appendage cobalt-coordinated (in a ‘‘base-on’’ form), or decoordinated, i.e., in a ‘‘base-off’’ form (see Figure 4) [22]. Depending upon their availability in either ‘‘base-off’’ or ‘‘base-on’’ forms, the reactivity of B12 derivatives in biologically relevant organometallic reactions is modified (as a consequence of the coordinating nucleotide function), as well as the face-selectivity at the corrin-bound cobalt center [104,125]: The DMB nucleotide function may be effective in directing alkylation (and other ligation) reactions (in cobalamins) to the ‘‘upper’’ (or b-face) in a thermodynamic sense, and it may stabilize the (organo)-B12 derivative in its ‘‘baseon’’ form [126,127]. As discussed below, this is particularly relevant in methyl-corrinoids, such as MeCbl (3) [126]. ‘‘Base-on’’ and ‘‘base-off’’ forms of the ‘‘complete’’ corrinoids represent constitutional isomers, that may (or may not) be well structured for binding by specific B12 apoenzymes: some B12-dependent enzymes, such as the methyltransferases, typically bind a ‘‘base-off’’ form of the B12 cofactor [36,128]. However, coenzyme B12-dependent enzymes come in two classes, Met. Ions Life Sci. 2009, 6, 1–51


17 R

R

+ Co

N

CH3

N

CH3

HO O

+ Co

in H2O

+ H2O

H 2O

OH O

O

O

−

O

−

O

O P

P

O

OH

“base-on”

− HX

+ HX

− HX

N

O OH

+ HX

N O

H3C

CH3

“base-off”

R

+ Co

H 2O

protonated “base-off”

−

X OH O −

O

+

O P

N

N

H

O O OH

H3C

CH3

Figure 4. B12 – a constitutional ‘‘molecular switch’’: The nucleotide base of B12 cofactors is cobalt-coordinated in the ‘‘base-on’’ form or de-coordinated in the ‘‘base-off’’ form; the two forms represent ‘‘constitutional’’ isomers and differ by their (bond) connectivity; protonation of the de-coordinated nucleotide base stabilizes the ‘‘complete’’ corrinoids in their ‘‘base-off’’ form.

one of them with a base-on B12 cofactor, the other with the B12 cofactor bound in a ‘‘base-off’’ (and ‘‘His-on’’) form (a still puzzling observation, see e.g., [129]). The unusual a-configuration of the ‘‘nucleotide moiety’’ is a common (stereochemical) feature of all known ‘‘complete’’ corrinoids. It allows, first Met. Ions Life Sci. 2009, 6, 1–51

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of all, for the intramolecular coordination of the heterocyclic nucleotide base to the ‘‘lower’’ a-axial coordination site of the corrin-bound cobalt center [11,12,32]. However, in some natural corrinoids non-coordinating phenols are found in their a-pseudonucleotide, such as p-cresol in p-cresolylcobamide (38) [11,130,131]. These remarkable findings may call for an additional ‘‘functional’’ rationalization of the differing properties of the ‘‘complete’’ corrinoids. The complete structure of the nucleotide moiety clearly is very important for the selective and tight binding by B12-binding proteins [132]. This is considered as the relevant basis for discriminating between the natural B12 derivatives by the human B12 uptake and transport system [16,133], which recognizes and binds its B12 load in the ‘‘base-on’’ form [132]. When binding ‘‘base-off analogues’’, such as pseudo-coenzyme B12 (34) or adenosyl-factor A (35), mammalian B12 transporters may even restructure them into their ‘‘baseon’’ constitution, which is the less stable form in aqueous solution [133]. A protein environment may thus bind and switch the bound B12 cofactors from ‘‘base-on’’ to ‘‘base-off’’ or vice versa [22,49]. In an artificially developed B12-binding protein, an antibody raised against coenzyme B12 (AdoCbl, 2), the bound analogues 34 and 35 were indeed found restructured into their base-on forms [134]. Similar phenomena may also arise in a(n oligo-) nucleotide environment [49]. Indeed, an interaction in the reverse sense was recently discovered in a particular type of mRNA, for which coenzyme B12 (AdoCbl, 2) is a strongly bound ligand: this part of mRNA in its untranslated region (a ‘‘B12 riboswitch’’) not only is able to bind coenzyme B12 (AdoCbl, 2) strongly, but it is also switched upon binding of the B12 ligand, so as to inhibit the expression of their gene product [135]. The cobalamins and related ‘‘complete’’ corrinoids thus have the capacity to switch their constitution between the ‘‘base-on’’ and the ‘‘base-off’’ forms and thus represent natural ‘‘molecular switches’’ [49]. The ‘‘base-on’’ to ‘‘baseoff’’ switch can also be achieved by protonation of the nucleotide base and decoordination from the corrin-bound cobalt ion [11,12]. The proton-assisted decoordination is inhibited by strong cobalt coordination, as is typical for 1 and other cyano-Co(III)-corrins, and is achieved readily in organic B12 derivatives, such as AdoCbl (2). The associated acidity of the protonated base-off form (as expressed by its pKa) thus informs quantitatively on the strength of the intramolecular coordination of the nucleotide base [11,12].

3. REDOX CHEMISTRY OF B12 DERIVATIVES Under physiological conditions vitamin B12 derivatives exist in three different oxidation states – Co(III), Co(II), and Co(I) – each possessing Met. Ions Life Sci. 2009, 6, 1–51


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different coordination properties and characteristically differing reactivities [5–12,22]. Oxidation-reduction processes are, therefore, of key importance in the chemistry and biology of B12. Electrochemical methods have been used for measuring the crucial redox potentials, for the purpose of generating reduced forms of protein-bound B12 derivatives [136] and electrode-bound B12 derivatives for analytical applications [137], as well as in the synthesis of organometallic B12 derivatives [138]. Axial coordination to the corrin-bound cobalt center depends on the formal oxidation state of the cobalt ion [11,12,138,139]: As a rule, the number of axial ligands decreases with the cobalt oxidation state. In the thermodynamically predominating forms of cobalt corrins, the diamagnetic Co(III) has two axial ligands bound (coordination number 6), the paramagnetic (low spin) Co(II) has one axial ligand bound (coordination number 5), and for the diamagnetic Co(I) no axial ligands are bound (coordination number 4), or only very weakly. Electron transfer reactions involving B12 derivatives are, therefore, accompanied by a change in the number of axial ligands, which, in reverse, heavily influence the thermodynamic and kinetic features of the electrochemistry of cobalt corrins [138,139]. In Co(III)-corrins, such as vitamin B12 (1) and hydroxocobalamin (5), the corrin-bound cobalt center binds two axial ligands (one of them the DMB base in the ‘‘base-on’’ cobalamins). In contrast, the metal center in Co(I)corrins, such as cob(I)alamin (39, B12s), is highly nucleophilic [140] and carries no axial ligand [139] (Figure 5). The intermediate oxidation state of Co(II)-corrins, such as in cob(II)alamin (6, B12r), provides a highly reactive metal-centered radicaloid species [58,141]. The use of electrochemistry thus provides an excellent means for generating, under controlled conditions, B12 derivatives of specific redox reactivity, as well as investigating the redox

R +

CoIII

+ e− −

+ e−

+CoII

DMB −O

(1, R = CN) vitamin B12 aquacobalamin (4+, R = H2O+)

CoI

− e−

e− −O

DMB

−O

cob(II)alamin (6)

DMB

cob(I)alamin (39−)

Figure 5. Outline of the redox transitions between cob(III)alamins (e.g. vitamin B12 (CNCbl, 1) or aquacobalamin (H2OCbl, 41)), cob(II)alamin (6), and cob(I)alamin (39). Met. Ions Life Sci. 2009, 6, 1–51

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processes in their interconversion between oxidation states, as studied and reviewed by Lexa and Saveánt [139].

3.1. Thermodynamics of Redox Processes The electrochemistry of the B12 derivative aquacobalamin (41) has been particularly well studied [139]: one-electron reduction of 41 first gives B12r (6) and then B12s (39) (see Figure 5). Typically, electrochemical studies of aquacobalamin (41) were carried out in aqueous solution. A complete standard potential versus pH diagram has been established that correlates the thermodynamics of the aquacobalamin (41)-B12r (6)-B12s (39–) system [139]. The interconversion between the different oxidation states of B12 derivatives can usually be seen with the eye or monitored effectively by UVVis spectroscopy. The redox data from potentiostatic measurements can thus be critically supported by UV-Vis spectroscopy [139,142]. Within the pH range 1 to 11 and applied potentials E ¼ 0.5 V and –1.2 V versus SCE, seven solution cobalamins predominate thermodynamically, spanning a range of the three formal oxidation states of B12 [138,139]. Aquacobalamin (41) and HOCbl (5) differ by protonation of the ‘‘upper’’ (b) axial ligand with pKa (41) ¼ 7.8 [139]. The Co(II)-corrin B12r (6) represents the ‘‘base-on’’ form of the Co(II) oxidation level (i.e., the nucleotide loop is coordinated intramolecularly), this is converted into the ‘‘base-off’’ (6-H1) by protonation of the DMB base, with pKa (6-H1) ¼ 2.9 [139]. At the Co(I) level, cob(I)alamin B12s (39–) is first protonated at the nucleotide base to give 39-H. For the pKa of 39-H, an original value of 4.7 was determined [139,143], but more recently this has been estimated to be 5.6 [144]. A second protonation then occurs at the Co(I) center to give the ‘‘Co(III)-hydride’’ + [145] 39-H+ 2 , with pKa (39-H2 ) ¼ 1 [139,146]. In the pH range 2.9 to 7.8, 41 and (base-on) B12r (6) represent the predominant Co(III)/Co(II) redox couple, with a standard potential of –0.04 V. For the Co(II)/Co(I) redox system there are two pH-independent standard potentials [139]: at a pH less than 5.6 the Co(II)/Co(I) couple (base-off) 6H1/39-H predominates at a standard potential of –0.74 V, but for the redox couple (base-on) B12r (6)/B12s (39) a more negative standard potential of 0.85 V [139] is required. This shift by about 110–140 mV to a more negative potential for the reduction of (base-on) B12r (6) when compared to that of the protonated base-off form 6-H1, reflects the selective stabilization of the Co(II)-corrin 6 by intramolecular nucleotide coordination [126,139]. A dependence of the standard potentials of the Co(III)/Co(II) redox couples occurs at approximately 60 mV per pH unit, at pH 47.8 for HOCbl (5)/B12r (6) and at pH Met. Ions Life Sci. 2009, 6, 1–51


21

o2.9 for 41/6-H1 and this reflects the effect of the removal by protonation of one axial ligand. An analogous dependence of the potential occurs between pH 2.9 and ca. 5.6 for 6/6-H1 as well as below pH 1 for the Co(II)/ Co(I) redox couple 6-H1/39-H+ 2 . At all pH values the disproportionation of Co(II)-corrins to Co(III)- and Co(I)-corrins is thermodynamically disfavored (the disproportionation equilibrium constant is below 1010) [139]. A complex interplay between the thermodynamic and kinetic factors of electron transfer reactions occurs in the analogous studies of vitamin B12 (1), due to the strongly coordinating cyano ligand [139]. Coordination of (one or two) cyanide ligands to the Co(III) center stabilizes it against reduction and the Co(III)/Co(II) standard potentials are shifted to more negative values [139,147]. Cyanide ions transform 1 into (base-off) dicyano-cob(III)alamin (1-CN) with an equilibrium constant of about 104 M1 [147]. Electrochemical studies of the ‘‘incomplete’’ diaquacobinamide (4021) (Figure 6) gave a standard electrochemical potential for the diaquacob(III)inamide (4021)/aqua-cob(II)inamide (411) couple of +0.27 V [139,148]. This corresponds to the extrapolated value for the highly acidic protonated base-off form (4-H21) of aquacob(III)alamin (41), with pKa (4H21) ¼ ca. –2.4 [139]. The potential of the corresponding aqua-cob(II)inamide (411)/cob(I)inamide (42) couple was determined as 0.73 V [139]. The standard potential of the redox couple between 411 and 42 is thus indistinguishable from that of the base-off cobalamins B12r-H1(6-H1)/B12s-H (39-H). Electrochemical studies of organometallic B12 derivatives are complicated due to the rapid and irreversible loss of the organic ligand upon reduction [139]. Low temperature conditions are therefore required to obtain pertinent thermodynamic data of organometallic B12 derivatives [149]. The standard

H2O

H2O 2+

CoIII

+ e− −

+

CoII

e−

+ e− −

CoI

e−

H2O O

O

NH

O

NH H

H

H CH3

CH3 OH

OH 402+

NH CH3 OH

41+

42

Figure 6. Outline of the redox transitions between cob(III)inamide (4021), cob(II)inamide (411) and cob(I)inamide (42). Met. Ions Life Sci. 2009, 6, 1–51

22

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potential (at –30 1C) for the methylcob(III)alamin (3)/methylcob(II)alamin redox couple was estimated as 1.60 V versus SCE [139,149] similar to the value obtained for the coenzyme B12/5 0 -deoxy-5 0 -adenosyl-cob(II)-alamin pair [144]. The standard potential of the typical Co(III)/Co(II) redox pair of organometallic B12 derivatives is significantly more negative than that of B12r (6)/B12s (39) and out of the reach of biological reductants. However, upon one-electron reduction of 3 fast decoordination of the nucleotide base occurs, followed by rapid decomposition to a methyl radical and cob(I)alamin (39). The thermodynamic features of B12-redox systems can be summarized as: (i) Intramolecular coordination of the nucleotide base or strong coordinating or nucleophilic ligands (such as cyanide ions) stabilize the corrin-bound cobalt center against one-electron reduction and shift the Co(III)/Co(II) redox couples to more negative potentials. (ii) The one-electron reduction of organometallic Co(III)-corrins typically occurs at more negative potentials than the Co(II)/Co(I) redox couple B12r/B12s [139]. Exceptions to this are provided by organometallic B12 derivatives with electron withdrawing substituents on the organometallic group, such as methoxycarbonylmethyl-cob(III)alamin [150].

3.2. Kinetics of the Redox Processes One-electron transfer reactions of cobalt corrinoids are accompanied by either cleavage or formation of a bond to an axial ligand. Typically, a reduction is accompanied by an expulsion, and an oxidation by the coordination, of the ligand [139]. The electron transfer step accordingly takes place either in a concerted fashion or in a rapid associated step with coordination or dissociation of the axial ligand. Electron transfer in the protonated Co(II)/Co(I) couple B12r-H1 (6-H1)/ 40.1 cm s1) as the presumed B12s-H (39-H) is fast in aqueous solution (kapp s axial water ligand is only kinetically weakly bound in the base-off Co(II)corrin 6-H1 [139,149]. However, when the aqua ligand in 6-H1 is substituted by a stronger axial ligand, e.g., by the nucleotide base as in base-on B12r, the electron transfer is slowed down sufficiently so that its kinetics can be conveniently measured by cyclic voltammetry [139,151,152]. For example, in 1 [139], the electron the Co(II)/Co(I) redox couple 6/39 kapp s ¼ 0.0002 cm s transfer is at least a thousand times slower than in the base-off forms 6-H1/ 39-H. Met. Ions Life Sci. 2009, 6, 1–51


23

The trend in kinetics for Co(III)/Co(II) couples follows the same trend as those for the corresponding Co(II)-/Co(I)-couples, albeit much slower. The Co(III)/Co(II) couple aquacob(III)alamin (41)/B12r (6) has a rate constant for heterogeneous electron transfer of about 10–5 cm s1 [139]. The electron transfer steps for the cyano-cob(III)- and cyano-cob(II)alamins 1-CN and 23-CN are slower still [139,147]. There is an approximate linear correlation between the equilibrium constant for the coordination of the axial ligand and the standard apparent rate constant for electron transfer [139]. This correlation has been rationalized by a model, in which stretching of the bond between the cobalt ion and the axial ligand represents the main factor of the kinetics of the electron transfer. As a consequence, kinetic and thermodynamic dependence of the electron transfer on the strength of the complexation of the axial ligands both add up, resulting in more negative reduction potentials as the strength of the ligand increases. Organocobalamins, such as coenzyme B12 (2) and MeCbl (3), have a different kinetic behavior from CNCbl (1) and other Co(III)-corrins with strong axial ligands [139,153,154]. Whereas the Co(III)/Co(II) reduction potentials are quite negative, the kinetics of electron transfer are fast. The one-electron reduction of 3 to the unstable methylcob(II)alamin anion (43) was estimated to have a rate constant of 1200 s1 at 30 1C. However, the product of the one-electron reduction of methylcobinamide (441), methylcob(II)inamide (45), has a half life of only about 0.1 s at 20 1C and decomposes into a methyl radical and cob(I)inamide (42) (see Figure 7). An Arrhenius plot of the kinetics of the decomposition of 45 gave the activation

CH3

CH3 +CoIII

CH3 + e−

CoII

− e−

CoI

H2O O

NH

O

H

NH

CH3 OH 44

O

H

NH

H

CH3 OH 45

CH3 OH 42

Figure 7. One-electron reduction of methylcob(III)inamide (441) gives methylcob(II)inamide (45), which rapidly decomposes into cob(I)inamide (42) and a methyl radical. Met. Ions Life Sci. 2009, 6, 1–51

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24

energy to be 19 kcal/mol and a pre-exponential factor A ¼ 1017.6 s1 [153]. From the values of the (Co–C) bond dissociation energy (37 kcal/mol) of MeCbl (3) [155] and the kinetics of the decomposition of the intermediate 43, the one-electron reduction is suggested to reduce the strength of the (Co–C) bond of 3 (by about 12 kcal/mol) to ‘‘half’’ of its value [139,155].

4. REACTIVITY OF B12 DERIVATIVES IN ORGANOMETALLIC REACTIONS Formation and cleavage of the (Co–C) bond in organometallic B12 cofactors are crucial steps not only in the reactions catalyzed by B12-dependent enzymes, but also for the chemistry of B12 derivatives in solution [9– 12,22,25,156–159]. The reactivity of B12 derivatives in organometallic reactions thus holds the key to the understanding of the biological activity of the B12-dependent enzymes. The most common methods for the preparation of such organometallic B12 derivatives typically rely on the efficient alkylation of Co(I)-corrins. One practical method is an electrochemical approach, as also described below. In solution cleavage and formation of the (Co–C) bond have been observed to occur in all three basic oxidation levels of the corrin-bound cobalt center [22,156–159]. So far, two main paths for these organometallic reactions have also been found to be relevant in enzymes: (i) the homolytic mode is typical of the reactivity of coenzyme B12 (AdoCbl, 2): 50 -adenosyl-CoðIIIÞ-corrin Ð CoðIIÞ-corrin þ 50 -adenosyl radical Formally it is a one-electron reduction/oxidation of the corrin-bound cobalt center and it results in the cleavage or formation of a single axial bond [11,22,158–160]. (ii) the nucleophile-induced, heterolytic mode is typical of the reactivity of MeCbl (3): methyl-CoðIIIÞ-corrin þ nucleophile Ð CoðIÞ-corrin þ methylating agent Formally, it involves a two-electron reduction/oxidation of the corrinbound cobalt center and the cleavage or formation of two (trans) axial bonds [11,22]. Met. Ions Life Sci. 2009, 6, 1–51


25

4.1. Formation of the (Co–C) Bond in Organocorrinoids One important type of reactivity of B12 derivatives is represented by the highly nucleophilic Co(I)-corrins [140]. These provide the basis of the standard (heterolytic) mode of formation of the (Co–C) bond, also important in methyl-corrinoids in enzyme-catalyzed methyl transfer reactions [23,161,162]. This mode is represented by the reaction of Co(I)-corrins with alkylating agents in the formation of the (Co–C) bond (and the nucleophileinduced demethylation of methyl Co(III)-corrins for the cleavage of the (Co–C) bond). Overall an oxidative trans addition occurs at the corrinbound cobalt center [125,163] (Figure 8). Alkylation at the corrin-bound Co(I) center normally proceeds via the ‘‘classical’’ bimolecular nucleophilic substitution (SN2) mechanism, where the Co(I)-corrin acts as a ‘‘supernucleophile’’ [140,164]. However, in certain cases alkylation occurs via a two-step one-electron transfer path, where Co(I)-corrins act as strong one-electron reducing agents and the process goes via Co(II)-corrin intermediates [112]. With ‘‘complete’’ corrins, such as B12s (39), either pathway results in alkylation at the b-face, which allows the nucleotide to coordinate at the a-face of alkylcobalamins, such as MeCbl [22,125]. When the nucleotide base has been changed from a DMB base to an imidazole, little effect on the thermodynamics of the methyl transfer reaction occurs [94]. The studies of Co(I)-corrins, like B12s (39), have shown the following reactivity patterns relevant for the SN2 alkylation pathway: (i) the nucleophilicity of Co(I)-corrins is virtually independent of the presence of the DMB nucleotide, both ‘‘complete’’ and ‘‘incomplete’’ Co(I)-corrins react preferentially at their b-face, which is essentially more nucleophilic [125]. The immediate product of the b-alkylation

CH3 CoI

SN2

+

+ CH3 X

CH3 Kon

CoIII

solv.

−

X −O

DMB

−

O

+CoIII

−O

DMB

X−

DMB

39−

3

Figure 8. Methylation of cob(I)alamin B12s (39 ) by an SN2 mode is directed to the ‘‘upper’’ b-face (by both, kinetic and thermodynamic reasons) and yields MeCbl (3). Met. Ions Life Sci. 2009, 6, 1–51

26

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may be a penta-coordinate (or already solvated and effectively hexacoordinate) Cob-alkyl-Co(III)-corrin; (ii) in aqueous solution and at room temperature the ‘‘base-on’’ (hexacoordinate) methylcob(III)alamin is more stable by about 4 kcal/mol than the ‘‘base-off’’ Coa-aqua-Cob-methylcob(III)alamin [126]. From NMR studies, the latter has been estimated to still be more stable in water, by around 7 kcal/mol, than the corresponding ‘‘base-off’’ and dehydrated form of Cob-methylcob(III)alamin, which has a pentacoordinate Cob-methyl-Co(III) center [165].

With ‘‘incomplete’’ cobalt-corrins the situation is again more complex, with two diastereoisomeric alkylation products often formed [11,12,112]. In specific cases, under suitable kinetic control, one of the alkyl-Co(III)-corrin diastereoisomers can form with high selectivity. For example with the Co(I) form of the lipophilic heptamethylcob(II)yrinate (11), the SN2-pathway can provide b-methylation with high diastereoselectivity (496%), whilst the one-electron transfer mechanism permits the formation of the Coa-methylated product with high diastereoselectivity (498%) [22,112]. However, configurational equilibration via rapid methyl group transfer reactions (involving Co(I)-, Co(II)-, and unalkylated Co(III)-corrins as methyl group acceptors) may give another overall outcome [22,126]. Electrochemistry is an excellent method for the selective and controlled production of reduced B12 forms under potentiostatic control. As alkyl halides or alkyl tosylates react quickly and efficiently with Co(I)-corrins [138], which are cleanly generated at controlled electrode potentials near that of Co(II)/Co(I) couples, electrochemistry provides a suitable method for the synthesis of organometallic B12 derivatives [139]. Indeed, the one-electron reduction of organometallic Co(III)-corrins typically occurs at more negative potentials than the Co(II)/Co(I) redox couple B12r/B12s [139]! Using electrolysis at a controlled potential of –1.1 V versus SCE, coenzyme B12 (2) was prepared in 95% yield from vitamin B12 (1) or from aquacobalamin (41) by alkylating cob(I)alamin (39) with 5 0 -chloro-5 0 deoxyadenosine [166]. Other organometallic B12 derivatives produced in an analogous method were, e.g., pseudocoenzyme B12 (37) (78% yield from pseudovitamin B12) [110], neocoenzyme B12 (39) (89% from neovitamin B12) [111] and homocoenzyme B12 (24) (99% from 41) [85]. Cob-methyl-imidazolylcobamide (31) (90% yield from Cob-cyano-imidazolylcobamide) [94] and methyl-13-epicob(III)alamin (46) (88% yield from neovitamin B12) [111] were synthesized by alkylation with methyl iodide. Also, dimeric B12 derivatives, such as the Cob-alkyl bridged and sterically crowded tetramethylene-Cob-1,4-biscobalamin (30) [98], and a strained organometallic B12-rotaxane [167], were synthesized by similar methods (see Figure 9). Met. Ions Life Sci. 2009, 6, 1–51

ORGANOMETALLIC CHEMISTRY OF B12 COENZYMES CH2

CH2)n

(CH2

27

CH2

H2NOC

H2NOC

CONH2

CONH2 CH3

H2NOC H3C H3C

N

N

CH3 H3C

H2NOC

CH3

N

N

O

H 3C

CH3 CONH2 O

HN

CH3

N

CH3 CH3

H3C

CONH2 HN

CONH2

Co+

H

CH3

CH3

N

N

H3C

Co+

H H2NOC

H 3C

CONH2

N

N

CH3

H2NOC

CH3

N

CH3

N

CH3

H 3C

H

HO O

N

CH3

H

HO O

O

O P

−

O

O

O P

O

OH

−

O

O

DMB

DMB

CoIII

CoIII

CoIII DMB

OH

CoIII DMB

Figure 9. Electrochemistry as means for the preparation of alkyl-bridged biscorrinoids. Structural formulae of tetramethylene-bridged biscobalamin (30, n ¼ 1) [97] and of a dodecamethylene-bridged biscobalamin (n ¼ 5); symbolic representations of alkyl-bridged biscobalamins and of a cyclodextrin-based B12-rotaxane [166]. Met. Ions Life Sci. 2009, 6, 1–51

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The high nucleophilicity of cob(I)alamin (39) towards alkylating agents makes it a versatile tool for the detection of toxicologically relevant electrophilic reagents. Such analytical methods are facilitating in vitro and in vivo studies of genotoxic compounds in cancer risk assessment. Many genotoxic compounds are directly (or indirectly) electrophilically reactive. The use of cob(I)alamin (39) as an analytical tool has been investigated in the trapping of oxiranes, metabolites of alkenes, to form alkyl-Cbls (Figure 10) [168,169]. It is presumed that the reaction proceeds according to an SN2 reaction following attack at the least hindered carbon [170]. In the work of Fred et al. [168], the 1,2-epoxide metabolites (oxiranes) of 1,3-butadiene were studied. For each metabolite a specific alkyl-Cbl was formed and it was possible to discriminate between the products by HPLCUV and by LC-MS. The cob(I)alamin (39), used in this study, had the advantage of reacting about 400,000 times faster than, e.g., nicotinamide, and therefore gave a better on-the-spot account [168]. Similar processes may also be relevant in vivo, as made likely by recent studies, which aimed to mimic the chemical reactions that could deplete vitamin B12 as a result of human exposure to electrophilic xenobiotics (styrene, chloroprene, and 1,3butadiene) [171]. The electrochemical methodology (see previous section) has been further developed as a method for the clean preparation of easily reduced

R′ H

H OH

H +

O

L +CoIII

+ 2 e-

CoI

−O

CoIII

DMB

R′ R′ H

−

O

DMB −

OH

H O

H

DMB +

−

CoIII

DMB O

Figure 10. Illustration of the alkylation of a cob(I)alamin (39–) by reaction with an oxirane [167]. Met. Ions Life Sci. 2009, 6, 1–51


29

organocorrinoids, such as Cob-[(methoxycarbonyl)methyl]-cob(III)alamin (47) via the alkylation of cob(II)alamin (6) [172]. Easily reducible organocob(III)alamins are known to be cleaved by direct electrochemical reduction or by reduction with cob(I)alamin (39) [150]. An acceptor-substituted C atom is directly bound to the Co center in 47 inducing it to be reduced at a peak potential of 0.90 V versus SCE (in DMF, room temperature) [173]. This value is close to that of the redox couple (base-on) B12r (6)/B12s (39) (0.85 V) and explains the difficulties encountered when preparing 47 via alkylation of 39– [173]. However, when aquacob(III)alamin chloride (41Cl) was submitted to a controlled potential of 0.45 V (versus 0.1 N CE) under potentiostatic conditions, it gave 6 cleanly. After the addition of an excess of methylbromoacetate and continued electrochemical reduction at 0.45 V, the crystalline alkyl cob(III)alamin was isolated in 75% yield [172]. The reaction was proposed to take place directly via 6 and radical intermediates [172]. The alkylation of ‘‘complete’’ Co(II)corrinoids with sufficiently reactive alkylating agents (methyl iodide, methylbromoacetate, etc.) thus is an efficient and alternative method to the more established synthetic procedures via Co(I)corrinoids for the synthesis of reduction-labile Co(III) organocorrinoids [172,174]. Organocobalamins are also accessible by the reaction between Co(II)corrins and radicals. In particular, the radicaloid cob(II)alamin (B12r, 6) has a penta-coordinated Co(II) center and can be considered to fulfill all the structural criteria of a highly efficient ‘‘radical trap’’, as revealed by the crystal structure of 6 [58]: the reactions of B12r (6) with alkyl radicals are indicated to occur with negligible restructuring of the (DMB nucleotide coordinated) cobalt corrin moiety and to furnish coenzyme B12 (2) and other organo-Cbls directly by the ‘‘homolytic’’ mode of formation of the (Co–C) bond [58]. From this it is understandable that the remarkably high reaction rate of 6 with alkyl radicals (such as the 5 0 -deoxy-5 0 -adenosyl radical), and the diastereospecificity for the reaction at the b-face, are both consistent and explainable due to the structure of cob(II)alamin (see Figure 11). The

R +

−O

CoII

Nu

CoIII

+

+ R

−O

Nu

6

Figure 11. The ‘‘radical trap’’ cob(II)alamin (6) rapidly combines with radicals on the ‘‘upper’’ b-face. Met. Ions Life Sci. 2009, 6, 1–51

KRAÜTLER

30

coordination of the DMB-nucleotide in 6 controls the (a/b)-diastereoface selectivity (in both a kinetic and thermodynamic sense) in alkylation reactions at the Co(II) center, which give b-alkyl-Cbls directly [141,159]. The stereochemical situation is appreciably more complex in ‘‘incomplete’’ corrins, such as cob(II)ester (11) and ‘‘base-off’’-forms of ‘‘complete’’ corrins [22]. The axial ligand at the corrin-bound Co(II) center is expected to direct the formation of the (Co–C) bond. In this way kinetic control can lead with high efficiency to the ‘‘rare’’ a-alkyl-Co(III)-corrins [112,175]. In such radical recombination reactions the axial ligand at the a- or b-side of the metal center will not only steer the diasteroselectivity of the alkylation but also may contribute to significant altering of the cage effects [160,176]. The two most relevant modes of formation (and cleavage) of the (Co–C) bond of the cobalt center differ significantly in their structural requirements (see Figure 12): The heterolytic mode of formation (and cleavage) of the (Co–C) bond, in which significant reorganizations at both faces of the corrin-bound cobalt center occur; The homolytic mode of formation (and cleavage), in which the cobaltcorrin portion of complete cob(III)amides (such as 2 and 3) hardly changes structure. Photolysis of methylcobalamin (MeCbl, 3) in deoxygenated aqueous solution saturated with (pressurized) carbon monoxide, gave acetyl-cobalamin in good yield and in a radical reaction, which was considered to finally involve the (re)combination of cob(II)alamin (11) with an acetyl radical [177]. This experiment turned out not to be relevant for the biological

+e− −e− R CoI

+ − CH3

+

CoIII

+R

+CoII

−R −O

Nu

supernucleophile

−O

methylating agent

Nu source of alkyl radical

−O

Nu

radical trap

Figure 12. Elementary reaction steps in organometallic and redox transformations of ‘‘complete’’ corrinoids, and their patterns of reactivity relevant for their cofactor function in B12-dependent enzymes. Met. Ions Life Sci. 2009, 6, 1–51


31

assembly of acetyl-CoA by the now well known enzyme acetyl-CoA synthase [178]. However, this organometallic transformation with a B12-derivative turned out to find considerable interest as a model for the ‘‘slaving-in’’ mechanism in radical reactions [179]. Another special mode of the formation of (Co–C) bonds in alkyl Co(III)corrins involves nucleophilic alkylating agents and the electrophilic properties of aqua-Co(III)-corrins [156,157,180]. A further means of preparing methyl-corrinoids is opened by methyl group transfer reactions between corrinoids and methyl-corrinoids and some other methyl-organic compounds [22,126,181].

4.2. Cleavage of the (Co–C) Bond in Organocorrinoids As coenzyme B12 (AdoCbl, 2) is considered to be a ‘‘reversible carrier of an alkyl radical’’ (or a reversibly functioning ‘‘radical source’’ [159]), the homolytic mode of the cleavage of the (Co–C) bond of 2 is of particular importance in its role as a cofactor. The strength of the (Co–C) bond of AdoCbl has been calculated to be about 30 kcal/mol by using detailed kinetic analyses of the thermal decomposition of 2 [159,160,182]. Considerable cage effects, and the presence of both ‘‘base-on’’ and ‘‘base-off’’ forms of 2, caused complications in the quantitative treatment of the homolytic (Co–C) bond dissociation energy (BDE) [160]. In several organocobalamins, the nucleotide coordinated ‘‘base-on’’ forms decomposed faster than their corresponding nucleotide-deficient organocobinamides or their protonated (‘‘base-off’’) forms of the organocobalamins [183,184]. The intramolecular coordination of the nucleotide was therefore considered to cause a ‘‘mechanochemical’’ means of labilizing the (Co–C) bond of organometallic B12 derivatives [159,183,184]. The extension of this idea to the enzymatic reactions with 2 as cofactor was disputed [58]. In the time since, crystallographic studies of coenzyme B12-dependent enzymes also helped to dismiss much of the original idea concerning the direct ‘‘mechanochemical’’ mechanism. They rather suggested the specific stabilization of the homolysis fragments to be an important means of producing destabilization in the protein-bound AdoCbl (2) – and thus activating 2 towards homolysis [58]. From the more recent crystal structures of AdoCbl-dependent enzymes, a distant aden(os)ine-binding pocket is now recognized to provide the required structural means for this [58,86,88,185]. Indeed, the contribution of the nucleotide coordination to the ease of homolytic cleavage of AdoCbl (2) was found to be relatively small: On the basis of available thermodynamic data concerning the coordination of the nucleotide in 2 and of the homolysis product cob(II)alamin (6), the Met. Ions Life Sci. 2009, 6, 1–51

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32

coordination of the nucleotide was estimated to weaken the (Co–C) bond by only 0.7 kcal/mol [22,126]. In contrast, in MeCbl (3) the intramolecular coordination of the nucleotide was determined to increase the homolytic (Co–C)-BDE of 3, by a bout 4 kcal/mol according to studies of the methylgroup transfer equilibrium between MeCbl (3)/cob(II)inamide (41) and methylcobinamide (44)/cob(II)alamin (6) [22], see Figure 13. The nucleophile-induced dealkylations of alkyl-Co(III)-corrins is another well known means for cleavage of the (Co–C) bond, in particular for methylCo(III)-corrins. It has been less studied with MeCbl (3), due to the impediment of the nucleophile-induced demethylation by the intramolecular coordination of the nucleotide base [125,186]. Indeed, thiolates demethylate methylcobinamide (44) to cob(I)inamide (42) approximately 1000 times faster than MeCbl (3) to B12s (39) [186], reflecting the strong stabilizing effect of the coordinated nucleotide in 3 [22,126]. This effect is of relevance also for enzymatic methyl group transfer reactions involving protein-bound Co(I)- and methyl-Co(III)-corrins, where considerable axial base effects are seen [187]. The electrophile-induced dealkylation of the cobalt-bound methyl group by polarizable metal ions, such as Hg21 ions, is a crucial path to methylmetal complexes, such as the poisonous Hg-CH3 ion [26]. Aqua-Co(III)corrins can also demethylate methyl-Co(III)-corrins slowly at room temperature [181].The coordination of the DMB nucleotide modifies the reactivity of the cobalt center by enhancing the ease of abstraction by electrophiles, in both a kinetic and thermodynamic sense [125]. The (Co–C) bond of alkyl-Co(III)-corrins is rather inert against proteolytic cleavage under physiological conditions. The acid-induced heterolytic cleavage of the (Co–C) bond of MeCbl (3) is not documented, the cleavage of coenzyme B12

H2O

CH3 +CoIII

−O

+

CH3 +CoIII

+CoII

Nu

Nu O

NH

O

H CH3

OH MeCbl (3)

Cbi(II) (41+)

NH

+CoII

+

−

Nu O

H CH3

OH MeCbi (44+)

B12r (6)

Figure 13. Methyl transfer reaction involving MeCbl (3) and methyl-cob(III)inamide (441) as methyl group donors and B12r (6) and cob(II)inamide (411) as methyl group acceptors. Met. Ions Life Sci. 2009, 6, 1–51

ORGANOMETALLIC CHEMISTRY OF B12 COENZYMES R

R

C

H H C

H

H

+CoIII

−O

33

H H

+CoII

Nu −O CO2CH2CH3

H3CH2CO2C R=

Nu

C H2C

CH3

Figure 14. Methylcobalamin (3) as a methylating agent for organic radicals, a hypothetical new mechanism of biological methylation [189].

(2) occurs in acidic aqueous solution at low pH [188], but less readily when compared to 2 0 -deoxycoenzyme B12 (48) and 2 0 ,3 0 -di-deoxycoenzyme B12 (49) [163]. The reactivity difference can be traced back largely to the effect of the ease of protonation of the cobalt-bound organic group [163]. Interestingly, the replacement of the DMB base by an imidazole in Cob-adenosylimidazolyl-cobamide also results in a more readily dealkylated analogue of AdoCbl [188]. A recently recognized further mode of cleavage of the (Co–C) bond of organometallic B12 derivatives, is represented by the radical-induced substitution at the cobalt-bound carbon center [22,98,189] (Figure 14). This type of thermodynamically favorable reaction holds strong interest due to the observation of unusual biological (C–C) bond forming reactions and methylations at seemingly inactivated carbon centers [190,191]. In a formally related radical abstraction reaction, the cobalt-bound methyl group of methylcobalamin (3) and other methylcorrinoids is rapidly abstracted by Co(II)-corrinoids, such as cob(II)inamide (411), (giving methylcob(III)inamide, 441) and cob(II)alamin (B12r, 6) (Figure 14) [22]. This type of reaction does not involve free methyl radicals and, under appropriate conditions (aprotic solvents), it is not (even) sensitive to the presence of molecular oxygen [192]. The (Co–C) bond of various organocorrinoids is cleaved homolytically by absorption of visible light and organometallic B12 derivatives have long been know to be sensitive to visible light [193], which induces cleavage with a Met. Ions Life Sci. 2009, 6, 1–51

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quantum yield of about 0.3 [141,194,195]. Organo-cob(III)amides are also labile to strong one-electron reducing agents, as it has been found that after one electron reduction of organyl-Co(III)-corrins the (Co–C) bond is considerably weakened [139,155]. As noted above, this aspect may render it difficult to prepare organo-cob(III)amides with electron-withdrawing substituents, via alkylation of the strongly reducing cob(I)amides [173].

5. ORGANOMETALLIC B12 DERIVATIVES AS COFACTORS AND INTERMEDIATES IN ENZYMES ‘‘Complete’’ methyl- and adenosyl-corrinoids, such as MeCbl (3) and AdoCbl (2), typically are considered to be the relevant cofactor forms of corrinoids in enzymatic reactions. These organometallic corrinoids are frequently observable in the resting states (and may be found in isolation forms) of functioning enzymes, and may then be bound characteristically ‘‘base-on’’, ‘‘base-off/His-on’’ or ‘‘base-off’’ to the protein part of the enzyme [18–23]. The catalytically equally important dealkylated cofactor forms, such as the Co(II)- and Co(I)-corrinoids cob(II)alamin (6) and cob(I)alamin (39), are less well observable species, and transient in the enzyme reactions, for reasons of their thermodynamic instability under typical physiological conditions. In most organisms, physiologically inactive forms of the (‘‘complete’’) corrinoids are taken up and converted into active organometallic cofactor forms enzymatically. In the human metabolism, (inactive) vitamin B12 (1) is converted into the adenosyl-corrinoid coenzyme B12 (2) by an ATP-using adenosyl-transferase, or into the methyl-corrinoid MeCbl (3) by methylation with S-adenosyl-methionine (SAM) in the methionine synthase complex [17]. However, in the course of the biosynthesis of ‘‘complete’’ corrinoids in microorganisms, such as of the cobalamins, ‘‘incomplete’’ organometallic B12 derivatives already play an important role at an early stage as obligate biosynthesis intermediates [196]: Thus, the biosynthetic build-up of the ‘‘complete’’ corrinoids firsts leads to the ‘‘incomplete’’ (organometallic) Cob5 0 -deoxy-5 0 -adenosyl-cobinamide (32), to proceed further to the ‘‘complete’’ B12 derivatives by assembly of the nucleotide moiety [196]. Organometallic B12 derivatives are also considered as intermediates in B12-dependent reductive dehalogenases [97], which play an important role in the detoxification of chlorinated compounds [197,198]. Several B12dependent dehalogenases have been purified with nearly all containing one or more iron-sulfur clusters, in addition to the corrinoid cofactor [97,197]. In the anaerobic bacterium Sulfurospirillum multivorans, which catalyzes the reductive dehalogenation of tetrachloroethene and Met. Ions Life Sci. 2009, 6, 1–51


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trichloroethene to cis-1,2-dichloroethene [97], a novel corrinoid cofactor was found that had slightly different catalytic properties to other cobamides [66]. This cofactor was isolated as norpseudovitamin B12 (15, Cob-cyano-700 adeninyl-176-norcobinamide or 176-norpseudovitamin B12), a homologue of pseudovitamin B12 (16) lacking the methyl group attached to carbon 176 [66]. In the B12-catalyzed reductive dechlorination of tetrachloroethylene the first step is likely to involve an electron transfer from the fully reduced Co(I)corrin (such as 39) to tetrachloroethylene, leading to a Co(II)corrin (such as 6) and the formation of a trichlorovinyl radical by loss of chloride [199]. Chlorovinylcobalamin (29) and vinylcobalamin (28) were thus synthesized as model compounds [200]. It was shown that chlorinated organometallic derivatives could be possible intermediates in reductive dehalogenation reactions, as 39 promoted reactions can reduce such compounds back to the active form of the catalyst [95,96].

5.1. Methylcorrinoids in B12-Dependent Methyltransferases The B12-dependent methyltransferases play an important role in amino acid metabolism in many organisms (including humans) as well as in one-carbon metabolism and CO2 fixation in anaerobic microbes [21]. The reactivity of the ‘‘supernucleophilic’’ Co(I)-corrins and of methyl-Co(III)-corrins make B12 derivatives ideal as cofactors in such enzymatic methyl group transfer reactions [11,12]. B12-dependent methionine synthase has been particularly well studied (see e.g. [21,201]) as have methyltransferases in aerobic acetogenesis (see, e.g., [202]), methanogenesis (see, e.g., [203]), and in the anaerobic catabolism of acetic acid to methane and CO2 (see, e.g., [204]). Various substrates act as sources of methyl groups, such as methanol, methyl amines, aromatic methyl esters, methylated heavy metals or N5methyltetrahydro-pterins (such as N5-methyltetrahydromethanopterin or N5-methyltetrahydrofolate). For N5-methyltetrahydrofolate as a source of the methyl group it has been suggested that the methyl group donor is more likely to be the protonated form of N5-methyltetrahydrofolate [21]. Thiols are the methyl group acceptors in methionine synthesis (homocysteine) [21] and methanogenesis (coenzyme M) [23]. In the anaerobic biosynthesis of acetyl-coenzyme A from one-carbon precursors the methyl group acceptor is suggested to be the nickel center attached to the Fe/S cluster [205]. The methyl group transfers catalyzed by methionine synthase (MetH) from E. coli [21] and other B12-dependent methyltransferases are all indicated to proceed with an overall retention of configuration (i.e., consistent with two nucleophilic displacement steps, each with inversion of Met. Ions Life Sci. 2009, 6, 1–51

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configuration) [206,207]. These stereochemical findings exclude free methyl cations or radicals as intermediates, even though, in a formal sense, the methyl transfer reactions catalyzed by B12 enzymes involve (nucleophilicbound) methyl ‘‘cations’’ and heterolytic cleavage/formation of the (CoCH3) bond. The methyl group transfer thus, relies on the catalytic properties of enzyme-bound Co(I)corrins and methyl-Co(III)-corrins [22] and is amenable to considerable control from the protein environment [21], due to the great structural changes expected to accompany the transitions from (tetracoordinate) Co(I)corrins to (hexacoordinate) methyl-Co(III)-corrins [11,12]. The X-ray crystal analysis of the B12-binding domain of MetH provided the first insight into the three-dimensional structure of a B12-binding protein [36,116,208,209]. The astounding revelation of this work was the finding that the cobalt-coordinating DMB nucleotide tail of the protein-bound cofactor MeCbl (3) was displaced by a histidine imidazole and extended into the core of the ‘‘Rossmann fold’’ [36,209]. Consequently, in methionine synthase the corrinoid cofactor is bound by histidine ligation to the metal center and in a ‘‘base-off’’ constitution, i.e., bound in a ‘‘base-off/His-on’’ mode. Various other B12-dependent methyltransferases are indicated to have either a ‘‘baseoff/His-on’’ bound methyl-Co(III)-corrinoid, or even a corrinoid cofactor in ‘‘base-off’’ form (where His-coordination is absent) [128] (Figure 15). In a catalytic cycle of B12-dependent methyltransferases the corrinoid is indicated to cycle between a methyl-Co(III)-corrin and a Co(I)-corrin [21,23]. The changing between the hexacoordinate methyl-Co(III) form and (presumably) tetracoordinate Co(I) form is accompanied by constitutional/ conformational changes which are highly likely to provide a means for controlling the organometallic reactivity of the bound cofactor [22], subject to H1 uptake or H1 release (see Figure 15). In response, a H1-mediated switch mechanism may result, mediated via the ‘‘regulatory’’ His-Asp-Ser triad, which provides the crucial conformational alterations associated with the enzyme [21,201,210]. The nucleophile-induced methyl group transfers, involving heterolytic cleavage and formation of the organometallic (Co– CH3) bond at the corrin-bound cobalt center, are expected to be in-line attacks (incoming nucleophile/CH3-group/leaving group) and to be subject to strict geometric control: a main role of the His-Asp-Ser-triad appears to be participating in maintaining conformational control of the mutual placement of the corrinoid cofactors and the enzyme-bound substrates [210,211]. A significant second role of the His-Asp-Ser triad in organometallic reactions is associated with the thermodynamic effect of the a-axial base coordination on the strength of the (Cob–CH3) bond. Solution studies showed a significant thermodynamic trans effect of the DMB coordination in methylcobalamin (3) [11,22,125] and of the imidazole coordination in Cob–CH3-imidazolyl-cobamide (27) [94] on heterolytic methyl group Met. Ions Life Sci. 2009, 6, 1–51


37 NH3+

NH+3 H

H3C

CO−2

S

homocysteine

S

CO−2

methionine CH3 CoI

+CoIII

N DMB

O

O−. . . . . . . . . . Enz

H

N DMB O

− ..........

O

NHAr

N

Enz

CH3

NHAr

N HN

HN H2N

H+

N

N H

tetrahydrofolate

H2N

N

N H

N5-methyl-tetrahydrofolate

Figure 15. Illustration of methionine formation catalyzed by MetH (Enz signifies the MetH-apoenzyme) [21], where the bound corrinoid shuttles between MeCbl (3), in a ‘‘base-off/His-on’’ form, and cob(I)alamin (B12s, 39).

transfer reactions. The result showed that the stronger coordinating (nitrogen) ligand stabilizes the methyl-Co(III) form against nucleophilic abstraction of the methyl group by about 4 kcal/mol in 3 [126]. This may be seen mainly as an ‘‘electronic’’ effect [11,22,125], consistent with the observation of anomalous structural trans effects in other methyl-Co(III) complexes [100]. More recent studies with 27 suggested the imidazole base exerts similar ‘‘electronic effects’’ as the DMB base in 3 but 27 is more basic and, therefore, imidazolyl-cobamides (or the ‘‘base-off/His-on’’ form of 3) are more readily protonated near neutral pH [94]. The His-Asp-Ser triad may then represent a ‘‘relay’’ for H1 uptake/release, assumed to function in the enzymatic methylation/demethylation cycles [212,213]. In conclusion, the axial (Co–Na) bond in the methyl-Co(III) form of the protein-bound cofactor of MetH (and other B12-dependent Met. Ions Life Sci. 2009, 6, 1–51

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methyltransferases) appears to have three important consequences. The weakening of this bond activates both (i) the methyl group for heterolytic abstraction by a nucleophile and (ii) the Co(II) form for reduction to the Co(I) form and (iii) helps to position the methyl-cob(III)amide cofactor for methyl group transfer [22,201,211].

5.2. Adenosylcorrinoids in Enzymes Dependent on Coenzyme B12 About ten coenzyme B12-dependent enzymes are now known. These enzymes are four carbon skeleton mutases, two diol dehydratases, ethanolamine ammonia lyase, two amino mutases and B12-dependent ribonucleotide reductase (see [18–20,214–216]). The coenzyme B12-dependent enzymes are disproportionately distributed in the living world. Only methylmalonylCoA mutase is indispensable in human metabolism. The coenzyme B12-dependent enzymes perform chemical transformations that are difficult to achieve by typical ‘‘organic reactions’’ [18]. With the exception of the enzymatic ribonucleotide reduction [215], the results of coenzyme B12-catalyzed enzymatic reactions correspond to isomerizations with vicinal exchange of a hydrogen atom and of a group with heavy atom centers. Homolytic cleavage of the (Co–C) bond of the protein-bound AdoCbl (2) to a 5 0 -deoxy-5 0 -adenosyl radical and cob(II)alamin (6) was indicated early to be the entry to H abstraction reactions induced by the 5 0 deoxy-5 0 -adenosyl radical [217]. Therefore, homolysis of the (Co–C) bond of 2, which is the thermally most easily achieved reaction of 2 in solution (homolytic Co–C BDE of about 30 kcal/mol [159,176]) appears to be its biologically most significant reactivity: coenzyme B12 (2) is characterized as a ‘‘reversible free radical carrier’’ [159]) (see Figures 11, 12, and 16). However, the homolysis of the (Co–C) bond of the protein-bound coenzyme needs to be accelerated by a factor of about 1012 to agree with the observed rates of reaction of catalysis by the coenzyme B12-dependent enzymes [159,160]. The deduced dramatic labilization of the bound organometallic cofactor towards homolysis of the (Co–C) bond is an intriguing feature of the coenzyme B12-dependent enzymes [159,160,215]. The mechanism of the enzyme- (and substrate-) induced labilization of the (Co– C) bond still is a much discussed problem. Covalent restructuring of the bound cofactor (except for the formation of the ‘‘base-off/His-on’’-form in the carbon skeleton mutases) is not indicated [86,216]. In addition, protein and solvent molecules can only weakly stabilize a radical center [218]. Steric distortions of the protein-bound cofactor are thus considered as a likely means for the enhanced rate of (Co–C) bond homolysis [85,86,88,216]. In Met. Ions Life Sci. 2009, 6, 1–51

ORGANOMETALLIC CHEMISTRY OF B12 COENZYMES HO

H2C

39

OH

O

Ade

+Co

+CoII

Nu

Nu

+ H 2C

−

O

coenzyme B12 (2)

OH

HO

O

Ade

−O

cob(II)alamin (6)

5′-deoxy-5′-adenosyl radical

Figure 16. Coenzyme B12 (AdoCbl, 2), a reversible source of the 5 0 -deoxy-5 0 -adenosyl radical and of cob(II)alamin (B12r, 6).

view of the available crystal structures of cob(II)alamin (6) [58] and of coenzyme B12-dependent enzymes [36–44], Halpern’s earlier suggestion of an ‘‘upwards conformational distortion’’ of the cobalt-corrin part of 2 [159] is not likely to be of relevance. However, labilization may come about largely from a protein- and substrate-induced strain on the organometallic group, separation of the largely nonstrained homolysis fragments and strong binding by the protein of the separated pair, 5 0 -deoxy-5 0 -adenosyl radical and 6 (in either a ‘‘base-off/His-on’’ or ‘‘base-on’’ form) [85,86,88,216]. One explanation is the existence in some of these enzymes of a binding interface (e.g., of an ‘‘adenosine-binding pocket’’) which does not allow for unstrained binding of the organometallic moiety [85,86,88,216]. Fixed placement of the corrin moiety at the interfaces of the B12-binding and substrate-binding/ activating domains appears to be of high significance and movements of the corrin moiety are not required. The ‘‘regulatory triads’’ logically appears not to be involved in proton-transfer steps and may conserve its structure largely during enzymatic turnover. ‘‘Electronic effects’’ of the axial trans ligand on the (Co–C) bond homolysis in 2 and MeCbl (3) are now seen to be of less importance [22]. To conclude, all coenzyme B12-dependent enzymes appear to rely on the reactivity of bound organic radicals, which are formed (directly or indirectly) by a H atom abstraction by the 5 0 -deoxy-5 0 -adenosyl radical, that originates from the homolysis of the (Co–C) bond of AdoCbl (2). In these enzymatic reactions, the 5 0 -deoxy-5 0 -adenosyl radical is the established reactive partner in the actual enzymatic reaction, so that 2 should be looked at as a ‘‘precatalyst’’ (or catalyst precursor) [22]. Coenzyme B12 (2) might then be considered to be a structurally highly sophisticated, reversible source for an Met. Ions Life Sci. 2009, 6, 1–51

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alkyl radical, whose (Co–C) bond is labilized in the protein bound state [159], and the main role of the bound cofactor AdoCbl (2) is indeed, the production and controlled presentation of the 5 0 -deoxy-5 0 -adenosyl radical from homolysis of its (Co–C) bond [159]. The function of the remaining Co(II)-corrin fragment 6 of the coenzyme (as a ‘‘spectator’’ or a ‘‘conductor’’) has recently again become a matter of discussion [129] and has been re-addressed by calculations [219]. The rearrangement steps of B12-dependent enzymatic rearrangements are now assumed to be accomplished by tightly protein-bound radicals that are controlled in their reaction space [18]. Consequently, the major functions of the enzyme concern not only the catalysis of its proper reactions but also the reversible generation of the radical intermediates and the protection of its proteinic environment from non-specific radical chemistry, called ‘‘negative catalysis’’ [217].

6. CONCLUDING REMARKS AND FUTURE DIRECTIONS The discovery of B12 coenzymes by Barker et al. [220] and of heir organometallic nature by Lenhert and Hodgkin [1], as well as subsequent studies of the organometallic chemistry and biological function of AdoCbl and MeCbl have helped to open the field of ‘‘bioorgano-metallic’’ chemistry. Clearly, Nature makes use of the capacities of organometallic catalysis in a remarkable way, as is particularly apparent, e.g., in alternative pathways of carbon fixation in anaerobes [205,221]. In the B12-dependent metabolism, the B12 cofactors are bound to proteins and are subjected to the mutual interaction with the proteins. Recently, natural B12-binding nucleotides have also been discovered and suggested to function as ‘‘riboswitches’’, relevant in a new form of controlling gene expression [222]. This finding has begun to open a new area of research in the B12 field [223,224] and to induce further studies and complementary work with B12 nucleotide conjugates [49,225]. Indeed, the capacity for mutual interaction between the evolved B12 cofactors and functional nucleotides is hardly explored. This subject may intensify the search for evidence for a role of corrinoids in an early form of life, such as is represented by the (hypothesis of an) RNA world [226]. Corrinoids clearly are unique compounds extending the capacity of (biological) organometallic catalysis. In another direction, recent studies on the proteins involved in the uptake of B12 derivatives in microorganisms [227], humans [133], and other Met. Ions Life Sci. 2009, 6, 1–51


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mammals are giving a good foundation for investigations on other complex questions, such as: (i) how corrinoids are selectively acquired from the environment, and what forms of mutual dependencies and symbiosis may result from the metabolic need of B12-catalyzed organometallic processes (see, e.g., [228]). (ii) how vitamin B12 derivatives can be used as carriers to shuttle small ligands or larger molecules into cells, simple and higher organisms [74], to diagnose and to influence their metabolism [73]. The growing understanding of the unique reactivity of corrinoids in organometallic processes may also lead to an increasing use of these natural cobalt complexes in the (in vitro and in vivo) analysis of normal and aberrant life processes, such as chemical modifications and damage to DNA [229]. Along the same lines, organometallic processes with B12 derivatives also provide a remarkable potential as the general basis of novel synthetic and analytical developments [137]. Clearly, the ‘‘most beautiful’’ cofactor [230] and its unique organometallic reactivity [22] will continue to fascinate not only the ‘‘B12 fraternity’’, but it will keep a special place in a range of current and future scientific developments.

ACKNOWLEDGMENTS Over the years our work was generously supported by the Austrian National Science Foundation (FWF P13595) and by the European Commission (HPRN-CT-2002-00195).

ABBREVIATIONS Ado AdoCbl BDE Cbi Cbl CE CNCbl CoA DMB

adenosyl 5 0 -deoxy-5 0 -adenosylcobalamin, adenosylcobalamin, coenzyme B12 bond dissociation energy cobinamide cobalamin (DMB-cobamide) calomel electrode cyanocobalamin coenzyme A 5,6-dimethylbenzimidazole Met. Ions Life Sci. 2009, 6, 1–51

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DMF ESR H2OCbl HOCbl HPLC LC-MS MeCbl MetH NHE NMR NOE Nu SAM SCE UV-Vis

dimethylformamide electron spin resonance aquacobalamin, B12a hydroxocobalamin high performance liquid chromatography liquid chromatography-mass spectrometry methylcobalamin methionine synthase normal hydrogen electrode nuclear magnetic resonance nuclear Overhauser effect nucleotide function S-adenosylmethionine saturated calomel electrode ultraviolet visible absorbance spectrum

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225. S. Gscho¨sser and B. Kraütler, Chem. Eur. J., 2008, 14, 3605–3619. 226. R. R. Breaker, in The RNA World, Ed. R. F. Gesteland, T. R. Cech and J. F. Atkins, , Laboratory Press, Cold Spring Harbor, 2006, p. 89–107. 227. C. Bradbeer, in Chemistry and Biochemistry of B12, Ed. R. Banerjee, John Wiley & Sons, New York, 1999, p. 489–506. 228. M. T. Croft, A. D. Lawrence, E. Raux-Deery, M. J. Warren and A. G. Smith, Nature, 2005, 438, 90–93. 229. J. Haglund, A. Rafiq, L. Ehrenberg, B. T. Golding and M. Tornqvist, Chem. Res. Toxicol., 2000, 13, 253–256. 230. J. Stubbe, Science, 1994, 266, 1663–1664.



2 Cobalamin- and Corrinoid-Dependent Enzymes Rowena G. Matthews Department of Biological Chemistry and Life Sciences Institute, University of Michigan, Ann Arbor MI 48109-2216, USA

ABSTRACT 1. INTRODUCTION. WHAT IS A CORRINOID? 2. CORRINOID-DEPENDENT METHYLTRANSFERASES 2.1. Overview of the Metabolic Roles of Corrinoid-Dependent Methyltransferases 2.2. Cobalamin-Dependent Methionine Synthase 2.3. Corrinoid-Dependent Methyltransferases in Methanosarcina spp. 2.4. Membrane-Associated Energy-Conserving Corrinoid Methyltransferase 2.5. The Corrinoid Iron-Sulfur Protein 2.6. Aromatic O-Demethylases 2.7. Reductive Dehalogenases 2.8. Modes of Activation of Corrinoid-Dependent Methyltransferases 2.9. Methyl Transfer in Fosfomycin Biosynthesis 3. ADENOSYLCOBALAMIN-DEPENDENT REARRANGEMENTS AND ELIMINATIONS 3.1. Enzymes Catalyzing Carbon Skeleton Rearrangements 3.1.1. Glutamate Mutase 3.1.2. Methyleneglutarate Mutase 3.1.3. Methylmalonyl-Coenzyme A Mutase

Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00053

54 54 56 56 60 67 72 72 76 77 79 82 84 86 86 89 90

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3.1.4. Isobutyryl-Coenzyme A Mutase 94 3.1.5. MeaA – A Mutase of Unknown Function 94 3.2. Aminomutases and Diol Dehydrases. Isomerization and Elimination 95 3.2.1. Diol Dehydrase 95 3.2.2. Ethanolamine Ammonia Lyase 100 3.2.3. Lysine 5,6-Aminomutase 101 3.2.4. Ornithine 4,5-Aminomutase 103 3.3. Adenosylcobalamin-Dependent Ribonucleotide Triphosphate Reductase 103 4. CONCLUDING REMARKS 106 ACKNOWLEDGMENTS 107 ABBREVIATIONS AND DEFINITIONS 107 REFERENCES 107 ABSTRACT: This chapter reviews the literature on cobalamin- and corrinoid-containing enzymes. These enzymes fall into two broad classes, those using methylcobalamin or related methylcorrinoids as prosthetic groups and catalyzing methyl transfer reactions, and those using adenosylcobalamin as the prosthetic group and catalyzing the generation of substrate radicals that in turn undergo rearrangements and/or eliminations. KEYWORDS: adenosylcobalamin methylcobalamin methyltransferase

1. INTRODUCTION. WHAT IS A CORRINOID? The structure of cobalamin, or dimethylbenzimidazolylcobamide, is shown in Figure 1. In cob(III)alamin derivatives like methyl- or adenosylcobalamin the cobalt is in the +3 oxidation state and is typically six-coordinate. Four nitrogens from the corrin macrocycle serve as the equatorial ligands, and a substituent of the corrin ring known as the nucleotide loop, which terminates in a dimethylbenzimidazole base, provides the lower axial or a ligand to the cobalt. Cobamides in which the benzimidazole moiety is coordinated to the cobalt are referred to as ‘‘base-on’’ cobamides. The upper axial or b ligand, shown as R in Figure 1, is a methyl group in methylcobalamin, an adenosyl group in adenosylcobalamin (AdoCbl), or may be occupied by an exchangeable ligand such as water in aquacobalamin. In the domains Archaea and Prokarya, cobalamin is only one of many variants grouped under the name corrinoids. Some of these variants involve changes in the structure of the dimethybenzimidazole (DMB) nucleotide base, such as 5 0 -methoxybenzimidazole cobamide, while other variants involve replacement of the DMB base by compounds such as adenine (pseudovitamin Met. Ions Life Sci. 2009, 6, 53–114

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Figure 1. The structure of cobalamin. R is a methyl group in methylcobalamin or an adenosyl group in adenosylcobalamin.

B12) or p-cresol (p-cresolylcobamide). The latter two bases cannot be coordinated to the cobalt of the free cobamide, which instead contains a water in the lower axial position and is referred to as a base-off corrinoid. Corrinoid-dependent methyltransferases are found in all three kingdoms of life, and in all cases, the cofactor is bound to its enzyme in a base-off manner. It is probably for this reason that so much variation in the nucleotide loop is tolerated. In a subset of corrinoid-dependent methyltransferases, the corrinoid is bound with a histidine (His) from the protein as Met. Ions Life Sci. 2009, 6, 53–114

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the lower axial ligand, and this mode of binding is referred to as base-off, His-on. The first observation of base-off binding was made by Ragsdale and his colleagues [1], who characterized the corrinoid in the corrinoid ironsulfur protein from Moorella thermoaceticum by electron paragmagnetic resonance (EPR) and Mo¨ssbauer spectroscopy. Extracts of the bacterium Sporomusa ovata were subsequently shown to contain the base-off corrinoid p-cresolyl-cob(II)amide in the Stupperich laboratory [2]. When a protein containing the bound corrinoid was isolated, the corrinoid was found to exhibit the EPR properties of a base-on corrinoid in the +2 oxidation state. When the bacterial cells were grown on [15N]-His, the EPR spectrum was altered, indicating that the axial nitrogen ligand of the corrinoid was derived from the imidazole group of His. However, after release from the protein, the corrinoid remained in the base-off form. AdoCbl-dependent enzymes are only found in the domains Eukaryota and Prokaryota, and cobalamin is the only corrinoid to be found in these enzymes. As will be discussed further in the second major section of this chapter, AdoCbl-dependent enzymes may contain the cobalamin bound either in the base-on form with the DMB ligand coordinated to the cobalt, or in the base-off,His-on form. As the cobalt in corrinoids is reduced, the preferred coordination number decreases. Corrinoids in the +2 oxidation state are preferentially fivecoordinate, with only one axial ligand, while corrinoids in the +1 oxidation state are preferentially four-coordinate, with no axial ligands.

2. CORRINOID-DEPENDENT METHYLTRANSFERASES 2.1. Overview of the Metabolic Roles of CorrinoidDependent Methyltransferases In humans, only one corrinoid-dependent methyltransferase, methionine synthase, is found, and this appears to be the only such corrinoid-dependent methyltransferase to be found in the domain Eukarya. However, in the domains Prokarya and Archaea, a wide variety of corrinoid-dependent methyltransferases play central roles in metabolism, particularly in organisms that grow under anaerobic conditions. We will begin by briefly enumerating these methyltransferases and their roles in carbon assimilation and energy generation. Many organisms belonging to these two domains make use of the reactions in the Wood-Ljungdahl pathway (Figure 2), first elucidated by the studies of Harland Wood and Lars Ljungdahl and their groups (recently reviewed in [3]). The enzymes in the Eastern branch of the Wood-Ljungdahl pathway catalyze the reduction of CO2 to form methyl groups that are Met. Ions Life Sci. 2009, 6, 53–114


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Figure 2. The role of corrinoid methyltransferases in the central metabolic pathways of anaerobic eubacteria and methanogens. The corrinoid methyltransferases are labeled in red. Red arrows indicate pathways, not shown in detail, that also involve corrinoid proteins. The boxed reactions are not part of the Wood-Ljungdahl pathway but are unique to methanogens. R 0 is tetrahydrofolate or a tetrahydropterin analogue of tetrahydrofolate. The charge on the nickel that serves as the methyl acceptor on acyl-CoA synthase/decarbonylase is shown as +1, but whether it is Ni11 or Ni0 remains a matter for debate [175,176].

intially bound to tetrahydrofolate or tetrahydropterin analogues of tetrahydrofolate. The reducing power that is needed is provided by three hydride ion transfers. In organisms that can grow on CO2 and hydrogen, hydrogenases catalyze the reversible conversion of hydrogen gas to hydride and a proton. The Western branch of the Wood-Ljungdahl pathway involves the reduction of CO2 to CO, catalyzed by CO dehydrogenase, and the incorporation of CO into a methyl-nickel bond to form an acetyl-Ni at the Ni-Ni metal center of acyl-CoA synthase. The acetyl group generated by carbonylation can then be transferred to coenzyme A (CoA) to form acetyl-CoA. While none of the reactions of the Eastern or Western branches of the Wood-Ljungdahl pathway involve corrinoids, the corrinoid iron-sulfur protein (highlighted in red in Figure 2) plays a central role in transferring the methyl group generated in the Eastern branch to the nickel in acyl-CoA synthase. All of the reactions in the Wood-Ljungdahl pathway are Met. Ions Life Sci. 2009, 6, 53–114

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reversible, and some organisms will run portions of the Wood-Ljungdahl pathway in reverse, as we shall see. A subgroup of organisms in the kingdom Archaea are obligate anaerobes that derive their carbon from CO2 when grown in the presence of hydrogen and produce methane as the final product. These organisms are known as methanogens. Methanogens convert a portion of the methyl groups generated in the Eastern branch of the Wood-Ljungdahl pathway into methane. This reaction occurs in two irreversible steps, in which the methyl group is first transferred to coenzyme M (ethanethiolsulfonate) by coenzyme M methyltransferases, and then reduced to methane by coenzyme M reductase. These two steps are energy generating, and are coupled to the creation of an ion gradient across the cell membrane that is used to generate energy for cellular growth. The energy-conserving coenzyme M methyltransferase in the methanogen Methanobacterium thermoautotrophicum is a complex containing a corrinoid protein. The complex catalyzes the transfer of a methyl group from methyltetrahydromethanopterin, a methyltetrahydrofolate analogue found in this organism, to the cobalt of the corrinoid protein, and thence to the sulfur of coenzyme M to form methylcoenzyme M. The basic pattern for corrinoid-dependent methyltransferases is shown in Figure 3. These enzymes comprise a minimum of three modules, a central module that binds the corrinoid and modules that present the methyl donor to the corrinoid in the cobalt(I) oxidation state, and activate the donor if necessary, and that present the methyl acceptor to the methylcorrinoid and activate it if necessary. These three modules may reside on three separate proteins, or they may be present as modules on a single polypeptide or on several polypeptides. A common feature of these methyltransferases is that they must stabilize both the methylcorrinoid and the corrinoid in the cobalt(I) oxidation state. Furthermore, they must be capable of undergoing

Figure 3.

Basic pattern for corrinoid methyltransferases.



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conformational changes that allow the corrinoid prosthetic group access to both donor and acceptor modules. Some methanogens can also grow on acetate by converting it to acetylCoA and then reversing the acyl-CoA synthase reaction to decarbonylate the acetyl group and produce CO and the methylated form of the corrinoid ironsulfur protein. The CO is oxidized to CO2 with the generation of a hydride ion. The reversal of the methyl transfers catalyzed by the corrinoid ironsulfur protein complex will then produce a methyltetrahydropterin, which will be converted to methane. The hydride needed for the conversion of methylcoenzyme M to methane is generated by the oxidation of CO. Members of the genus Methanosarcina can also use other simple onecarbon compounds as sources of carbon and energy, including methylamines, methylthiols, and methanol. The methyl groups of these compounds are transferred to coenzyme M by specific non-energy conserving corrinoid methyltransferases that follow the basic pattern shown in Figure 3. Methanogens growing on substrates other than acetate synthesize acetylCoA from methyltetrahydropterins and CO2 by reversal of the corrinoid iron-sulfur complex methyl transfers to generate methyl-Ni on acyl-CoA synthase, followed by the acyl-CoA synthase reaction and oxidation of the resultant CO. From the acetyl-CoA thus formed, all other carbon-containing cellular components are generated. Acetogenic bacteria do not generate energy by methanogenesis, but rather generate energy by the anaerobic fermentation of glucose or by anaerobic growth on hydrogen and CO2. Glucose is converted to two molecules of pyruvate, which in turn is decarboxylated to form two molecules of acetylCoA in a reaction coupled to the generation of ATP from ADP and inorganic phosphate. They use the Eastern branch of the Wood-Ljungdahl pathway to reduce CO2 to a methyl group and the Western branch of the pathway to produce CO. These two reagents are then coupled to form an additional molecule of acetyl-CoA, from which all other carbon-containing compounds are generated. A variant use of the Wood-Ljungdahl pathway is made by hydrogenogenic bacteria such as Carboxydothermus hydrogenoformans, which can grow on CO as the sole source of energy and carbon under anaerobic conditions. Some of the CO is oxidized to CO2 by the action of CO dehydrogenase, with protons serving as the terminal electron acceptors. The Eastern branch of the Wood-Ljungdahl pathway is used for production of methyl groups, using hydride equivalents generated by oxidation of CO to CO2. The remainder of the CO is converted to acetyl-CoA by the action of acyl-CoA synthase and the corrinoid iron-sulfur protein. Another variant use of the Wood-Ljungdahl pathway is provided by bacteria that use aromatic O-methyl ethers as the source of both carbon and energy such as Sporomusa ovata. Corrinoid aromatic O-demethylases Met. Ions Life Sci. 2009, 6, 53–114

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catalyze transfer of the methyl group to tetrahydrofolate. The methyl group can then be oxidized to formate and/or CO2 by reversal of the Eastern branch of the pathway with the accompanying generation of reducing equivalents, and then converted to acetate using corrinoid iron-sulfur protein and acyl-CoA synthase. Finally corrinoid-dependent reductive dehalogenases found in Prokaryota use the corrinoid protein to catalyze the anaerobic dehalogenation of a variety of halogenated alkyl and aryl compounds. Since the products of such dehalogenations will vary with the halogenated substrate, the reductive dehalogenases are not shown on the central metabolic scheme in Figure 2.

2.2. Cobalamin-Dependent Methionine Synthase Cobalamin-dependent methionine synthase (MetH) catalyzes the reaction shown in equation (1). O

CH3 N

HN

O +

R

NH3 +

H2N

N

N H

Methyltetrahydrofolate

HS

COO–

L-Homocysteine

HN

H N

R

+

NH3 + H3C

H2N

N

N H

Tetrahydrofolate

S

COO–

L-Methionine

ð1Þ The enzyme is found in many members of the kingdom Prokaryota, including Escherichia coli, but has not been found in the Archaea. It is one of only two B12-dependent enzymes found in humans and other mammals, and is widely distributed among the animal Eukaryota. Due to its overexpression in recombinant form [4,5] and the resultant ease of purification under aerobic conditions, large amounts of purified E. coli protein have been available for biochemical and structural characterization. It was one of the first corrinoid proteins to be characterized, and has subsequently been extensively studied in the laboratories of Herbert Weissbach, Frank Huennekens, and more recently in my own laboratory. The enzyme consists of four modules, that are arranged linearly with single interdomain linkers to form a single 136 kDa polypeptide. The N-terminal module, the methyl donor module in the parlance used in Figure 3, binds and activates methyltetrahydrofolate and presents it to the cobalamin prosthetic group for methyl transfer. The next module in the sequence is the methyl acceptor module, this module binds and activates homocysteine and presents it to methylcobalamin to allow methyl transfer to form methionine. The third module is the cobalamin-binding module and also contains a four helix bundle Met. Ions Life Sci. 2009, 6, 53–114


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at the N-terminus of the domain. The final module binds adenosylmethionine (AdoMet) and is required for reductive activation of the protein. Its raison d’eˆtre requires discussion of the reactions catalyzed by the enzyme, which are shown in Figure 4. MetH is active during aerobic growth in E. coli, and also under in vitro turnover in microaerophilic conditions. In vitro, the cob(I)alamin form of the enzyme is oxidized to the inactive species cob(II)alamin about once in every 2000 turnovers [6]. Return of the prosthetic group to the active methylcobalamin form requires a reductive remethylation. In E. coli, the electron is provided by the electron transfer protein flavodoxin, the fldA gene product [7,8]. The reduction potential of the flavodoxin semiquinone/hydroquinone is 440 mV vs. the standard hydrogen electrode [9], and the quinone/ semiquinone potential, which is probably the more relevant one for cells grown under microaerophilic conditions, is 260 mV. In contrast, the cob(II)alamin/

Figure 4. Reactions catalyzed by methionine synthase. During primary turnover, the enzyme-bound cobalamin cycles between methylcobalamin and cob(I)alamin forms as the prosthetic group is alternately methylated by methyltetrahydrofolate (CH3-H4folate) and demethylated by transfer of the methyl group to homocysteine (Hcy). During turnover under microaerophilic conditions, the cob(I)alamin form of the enzyme is oxidized to cob(II)alamin about once in every 2000 turnovers. This form of the enzyme is inactive, and reactivation requires a reductive methylation in which the reduction of cob(II)alamin to cob(I)alamin is coupled to a highly exergonic methylation using adenosylmethionine (AdoMet) as the methyl donor. Met. Ions Life Sci. 2009, 6, 53–114

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cob(I)alamin reduction potential is 490 mV at pH 7 [10]. Thus the reduction of cob(II)alamin is a highly endergonic reaction and must be driven to completion by coupling to a highly exergonic methyl transfer. For this purpose, AdoMet is used as the methyl donor for reductive activation. The transfer of the methyl group of AdoMet is associated with a driving force of about 17 kcal/mol, helping to assure that the reductive remethylation proceeds quantitatively. The C-terminal domain of MetH binds AdoMet and catalyzes this alternate methyl donor reaction, and is designated as the reactivation module. Indeed, if this module is removed from the protein by limited proteolysis of the native enzyme, the methylated enzyme turns over until all the cobalamin accumulates as cob(II)alamin, at which point the enzyme can no longer be reactivated [6]. Further insights into the complicated catalytic and reactivation cycles of methionine synthase came as X-ray structures were determined for fragments of methionine synthase in the laboratory of Martha Ludwig. The entire enzyme has never been crystallized, presumably because of the conformational lability of the enzyme. The first fragment to be crystallized was the cobalamin-binding module [11]. This was the first structure of cobalamin bound to a protein, and it revealed the remarkable displacement of the DMB axial ligand by a histidine residue from the protein--what we now call the base-off,His-on state of cobalamin. As shown in Figure 5, His759 is coordinated to the cobalt of methylcobalamin and also linked by a network of hydrogen bonds to the carboxyl group of Asp757, which in turn is hydrogenbonded to the hydroxyl of Ser810. The binding of the prosthetic group in the base-off,His-on form was shown to be associated with a signature His-xAsp-xxGly---41/42---Ser-x-Leu-25/26---Gly-Gly sequence that had originally been identified in glutamate mutase [12]. The b-face of the cobalamin prosthetic group in the structure is shielded by a four helix bundle that forms the N-terminal portion of the module sequence and is referred to as the ‘‘cap’’. Thus far, only indirect evidence suggests that this conformation of the intact MetH protein exists in solution [13]. The structure of the C-terminal reactivation module of MetH was determined next [14], and then a structure was obtained of the entire C-terminal half of the protein comprising the cobalamin-binding and reactivation modules [15]. This structure was determined with a fragment of His759Gly MetH, and revealed that the fragment had crystallized in a conformation in which the b-face of the cobalamin prosthetic group was now in contact with the reactivation module (Figure 6, right). Although, His759 is absent in this structure, the distance between Ca of Gly759 and the cobalt of the cob(II)alamin prosthetic group is 2.3 A˚ greater than in the Cap:Cob conformation assumed by the isolated cobalamin-binding module. This increased distance would be predicted to cause the cobalamin of the wild-type enzyme to assume a base-off,His-off conformation, enforced in part by the juxtaposition of residues from the AdoMet-binding module between the cobalamin-binding domain and Met. Ions Life Sci. 2009, 6, 53–114


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Figure 5. Coordination of methylcobalamin in the cobalamin-binding module of MetH. A hydrogen bonded network comprising His759, Asp757 and Ser810 links His759 to the external solvent; these three residues that are absolutely conserved in all MetH enzymes, are referred to as the ligand triad. Reprinted from [177] with permission of Annual Reviews of Biochemistry, copyright 1997.

the corrin ring. Indeed, the methylated form of the wild-type enzyme has been shown to undergo interconversions between His-on and His-off forms that are induced by binding of ligands, shifts in temperature, or the binding of flavodoxin [15,16]. These interconversions are thought to reflect rearrangements of the four modules of methionine synthase, as shown in Figure 7. The left hand side of Figure 6 shows the structure of the N-terminal substrate-binding modules of methionine synthase from Thermotoga maritima [17]. The homocysteine- and folate-binding modules of methionine synthase are both a8b8 barrels, with their openings positioned orthogonally with respect to each other. There is a large buried surface area between the two barrels, strongly suggesting that they move as a unit rather than individually. Thus, as cartooned in Figure 7, large modular rearrangements are required to allow the cobalamin to access the Hcy-binding and Fol-binding modules alternately during catalysis. Originally, it was proposed that the base-off,His-on state of cobalamin in methionine synthase might accelerate the methyl transfer reactions [11]. However, mounting evidence suggests that the primary role of the ligand Met. Ions Life Sci. 2009, 6, 53–114

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Figure 6. Structures of the N-terminal and C-terminal halves of methionine synthase. The structure of the homocysteine-binding (green) and folate-binding (gold) modules of MetH was determined with protein from Thermotoga maritima [17], and the structure of the cobalamin-binding (red) and reactivation (blue) modules of MetH was determined with the His759Gly mutant of the enzyme from Escherichia coli [178]. The cobalamin-binding module also contains a four helix bundle, referred to as the ‘‘cap’’ and shown in grey.

replacement is to facilitate the conformational changes necessary for catalysis. In the initial studies, the wild-type His759 enzyme was compared with mutations of each residue of the ligand triad: His759Gly, Asp757Glu and Asp757Asn, and Ser810Ala. While the His759Gly mutant was inactive in steady-state assays, the Asp757 mutants showed kcat values that were 4–6% of the wild-type enzyme and that for the Ser810Ala mutant was 56% [5]. When the approach to steady-state was examined by enzyme-monitored turnover, the Asp757 mutants were 33 to 54% as fast as the wild-type enzyme and the Ser810Ala mutant was 61% as fast, indicating that these mutants were barely compromised in establishing the initial steady-state distribution of methylcobalamin and cob(I)alamin enzyme forms. The rate constant for the AdoMet- and reduced flavodoxin-dependent reactivation of enzyme in the cob(II)alamin form, which occurs in state 4 of Figure 7, was also measured. The His759Gly mutant was methylated 14 times faster than wild-type enzyme, as was the Asp757Glu mutant. The Asp757Asn and Ser810Ala mutants were methylated about twice as fast as the wild-type enzyme. Based on these data, Jarrett [18] proposed that mutations of residues in the ligand triad might alter the distribution of states Met. Ions Life Sci. 2009, 6, 53–114


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Figure 7. Conformational states of methionine synthase. The four modules are shown in gold (Hcy-binding), green (folate-binding), red (cobalamin-binding domain) and gray (cap domain), and blue (AdoMet-binding). The corrin ring of methylcobalamin is indicated by the rectangle on top of the cobalamin-binding domain, and His759 is indicated by the vertical line. In the AdoMet:Cob conformation, the histidine is displaced as indicated in the cartoon, and the corrin ring tilts away from the cobalamin-binding domain and from Ca of His759. Reprinted from [20] with permission of the National Academy of Sciences of the USA.

shown in Figure 7, and that, as the ligation of the histidine was weakened and then finally abolished, the distribution would increasingly favor the AdoMet:Cob conformation. In agreement with this proposal, it was found that the EPR spectra of wild-type and mutant enzymes in the cob(II)alamin form increasingly favored the His-off conformation in the order: His759Gly (100%) 4 Asp757Glu (65%) 4 Asp757Asn (25%) 4 wild-type (15%) 4 Ser810Ala (5%). The next contribution to our understanding was the demonstration that addition of oxidized flavodoxin to methionine synthase in the cob(II)alamin form resulted in the conversion of the His-on state of the prosthetic group to the His-off state, as shown by the loss of superhyperfine coupling in the EPR spectrum of the cobalamin [9]. Further studies [19] established that enzyme in the presumably His-off four-coordinate cob(I)alamin form could not interconvert between catalytic conformations (states 1, 2, and 3 in Figure 7) and the AdoMet:Cob conformation (state 4 in Figure 7). If MetH in the cob(I)alamin state is produced by reduction of cob(II)alamin, the enzyme thus formed reacts with AdoMet but not with methyltetrahydrofolate, suggesting that the enzyme can assume the AdoMet:Cob conformation but not the Fol:Cob conformation. Conversely, if cob(I)alamin is generated by demethylation of enzyme in the methylcobalamin state in the presence of homocysteine, the cob(I)alamin reacts with methyltetrahydrofolate 30,000fold more rapidly than it reacts with AdoMet, suggesting that it can readily assume the Fol:Cob conformation but has very limited access to the AdoMet:Cob conformation. Further evidence that the two forms of cob(I)alamin are in different protein conformations came from the observation that limited proteolysis of the native enzyme resulted in different patterns. The Met. Ions Life Sci. 2009, 6, 53–114

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pattern seen with cob(I)alamin enzyme generated by reduction was also seen with wild-type cob(II)alamin enzyme with flavodoxin bound (previously shown to be His-off [9]) and also with cob(II)alamin bound to the His759Gly mutant. In contrast, the cob(I)alamin enzyme generated by demethylation with homocysteine showed the same cleavage pattern as wild-type enzyme in the methylcobalamin and cob(II)alamin (in the absence of flavodoxin) forms. Thus the results suggested that the first cleavage pattern was characteristic of enzyme in the AdoMet:Cob conformation, while the second cleavage pattern was characteristic of enzyme in one of the catalytic conformations (states 1, 2, and 3 in Figure 7). The results described thus far indicated that enzyme in the cob(I)alamin form can not freely interconvert between catalytic and reactivation conformations, while enzyme in the cob(II)alamin form interconverts readily on addition of flavodoxin. Bandarian et al. [15] discovered that enzyme in the methylcobalamin form can also be induced to interconvert between catalytic and reactivation conformations, and that the reactivation conformation is associated with an absorbance spectrum typical of base-off methylcobalamin. He showed that the His-off/His-on equilibrium was o5:95 for the full length wild-type enzyme, while it was 12:88 for the Asp757Glu mutant, confirming that this ligand triad mutation weakens the ligation of His759 to the cobalt. Addition of AdoHcy, which is bound at the interface between the AdoMet module and the Cob module in the AdoMet:Cob conformation of the protein, shifts the equilibrium to favor the His-off state, consistent with the argument that the His-off state is associated with the AdoMet:Cob conformation. Addition of methyltetrahydrofolate also shifts the equilibrium to favor the His-off state, presumably because the methyl group of methyltetrahydrofolate is in steric conflict with the methyl group of methylcobalamin in the Fol:Cob state, which is therefore disfavored. The two ligands exert their effects on the equilbrium independently, favoring the His-off state with free energy changes of 0.9 and 0.6 kcal/mol, respectively. The picture that emerges is of a delicately balanced equilibrium between alternate conformations of methionine synthase, with small free energy changes induced by ligand binding able to shift the distribution of conformers because these ligands have different affinities for the different states. The next advance in our understanding of the dynamic equilibrium of conformers in MetH came from the studies of Fleischhacker [16]. She examined the effect of substitutions in the upper axial (b) ligand of cob(III)alamin on the conformational equilibrium. She showed that methionine synthase in the propylcobalamin form had a His-off/His-on equilibrium of 31:69 in the absence of ligands, while the His-off form was undetectable when the enzyme prosthetic group was aquacobalamin. The His-off/His-on equilibrium was predicted by the ligand trans influence, with the more electron-donating propyl substituent favoring the AdoMet:Cob conformation. Met. Ions Life Sci. 2009, 6, 53–114


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These studies provided a rationale for the base-off,His-on substitution. The histidine ligand serves as a protein sensor of the oxidation and ligation state of the cobalamin, biasing the equilibrium in accord with the net formal charge on the cobalt and its resultant effect on the histidine ligation. The cob(II)alamin state of methionine synthase showed properties intermediate between methylcobalamin and propylcobalamin in its ability to enter the AdoMet:Cob conformation. Oxidation of the prosthetic group to the cob(II)alamin state would lead to an enhanced propensity to enter the AdoMet:Cob conformation, which is favored by addition of flavodoxin and/ or by addition of AdoMet (there is no steric conflict between cob(II)alamin and AdoMet). Once the prosthetic group is reduced and methylated, the resultant His-off methylcobalamin is converted to the His-on state and thus returns to the catalytic cycle. A further role for the His759 ligand was discovered when the structure of a C-terminal fragment of ‘‘wild-type’’ methylated enzyme was determined [20]. To stabilize the AdoMet:Cob conformation of the enzyme, a disulfide crosslink was introduced between the ‘‘cap’’ and the cobalamin-binding domain by mutation of Ile690 and Gly743 to cysteine residues. The crystal structure of this fragment revealed that it was indeed in the AdoMet:Cob conformation, but that the histidine had now moved about 7A˚ away from the cobalt and was now involved in intermodular hydrogen bonding with the AdoMet-binding domain (Figure 8). These unanticipated intermodular contacts would be expected to stabilize the His-off forms of the wild-type enzyme in the AdoMet:Cob conformation by as much as 3–5 kcal/mol.

2.3. Corrinoid-Dependent Methyltransferases in Methanosarcina spp. Methanogens in the genus Methanosarcina use protein complexes containing corrinoid-binding proteins to catabolize simple one-carbon compounds such as methylamines and methylthiols as well as methanol. These complexes typically consist of a substrate-specific methyltransferase, a cognate corrinoid protein that receives the methyl group, and a second methyltransferase that catalyzes the transfer of the methyl group from the corrinoid protein to coenzyme M (ethane thiol sulfonate). More recently, tetramethylammonium-coenzyme M methyltransferase activity has been identified in Methanococcoides sp. [21]. Figure 9 diagrams the individual complexes that have been studied: methanol-coenzyme M methyltransferase [22], dimethylsulfide-coenzyme M methyltransferase [23], monomethylamine-coenzyme M methyltransferase [24,25], dimethylaminecoenzyme M methyltransferase [26], trimethylamine-coenzyme M methyltransferase [27], and tetramethylammonium-coenzyme M methyltransferase [28]. However, the genome sequence of Methanosarcina acetivorans contains 10 Met. Ions Life Sci. 2009, 6, 53–114

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Figure 8. Intermodular contacts between His759 and residues in the AdoMetbinding reactivation module of MetH. Ne2 of His759 interacts with the AdoMet module directly through a hydrogen bond to Asp1093 and via a water-mediated hydrogen bond to Glu1069. Nd1 of His759 forms a hydrogen bond with the amide of the propionamide side chain of ring B of the cobalamin (not shown). Adapted from [20] with permission of the National Academy of Sciences, USA.

sequences with homologies to substrate-specific methyltransferases, 15 putative corrinoid protein sequences, and 14 sequences with homologies to coenzyme M methyltransferases. The substrates for many of these proteins remain unidentified [28, 29]. The reactions catalyzed by these cytoplasmic enzymes bear striking similarity to the overall reaction catalyzed by cobalamin-dependent methionine synthase, and indeed the corrinoid-binding proteins in those complexes that have been sequenced show homology with the cobalamin-binding domain of methionine synthase, including the characteristic Asp-X-His-X-X-Gly motif indicative of a corrinoid cofactor with a histidine axial ligand. However, the methylcorrinoid-coenzyme M methyltransferase proteins do not show significant sequence homology with the homocysteine-binding domain of methionine synthase, but instead show sequence similarity with uroporphyrinogen decarboxylase (UroD) [30]. The substrate-methylcorrinoid methyl Met. Ions Life Sci. 2009, 6, 53–114

COBALAMIN- AND CORRINOID-DEPENDENT ENZYMES MtaB

MtaC

MtaA

Co

methanol H2O MtsA

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CH3-SCoM

CoMSH MtsB

MtsA

Co

dimethylsulfide (methylmercaptopropionic acid)

CH3-SCoM

methanethiol CoMSH (mercaptopropionic acid) MtmB

MtmC

MtbA

Co

monomethylamine NH4+ MtbB1

CH3-SCoM

CoMSH MtbC

MtbA

Co

dimethylamine

monomethylamine

MttB1

MttC

CH3-SCoM CoMSH

MtbA

Co

trimethylamine dimethylamine

MtqB

MtqC

CH3-SCoM CoMSH

MtqA

Co

tetramethylammonium trimethylamine

CH3-SCoM CoMSH

Figure 9. Complexes catalyzing methyl transfer from methyl donors to coenzyme M. The gene product designations are shown above each protein. The first two letters, mt, indicate involvement of the gene product in methyl transfer, the third letter indicates the substrate: a for methanol, s for methylthiols, m for monomethylamine, b for dimethylamine, t for trimethylamine and q for tetramethylammonium. The final letter designates the polypeptide function: where B is the substrate-specific methyltransferase that methylates the corrinoid protein with a methyl group derived from substrate, C is the corrinoid binding polypeptide and A is the CoM-methylating protein, as originally suggested by Krzycki and coworkers [27].


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transferases neither resemble methionine synthase domains nor each other. The genes specifying MtmC, MtbC, and MttC all contain an in-frame UAG stop codon in the middle of the open reading frame [27], and this UAG has been shown to specify pyrrolysine [31]. This unique residue is located at the active site of MtmB and is thought to be involved in activation of the amine substrate for methyl transfer. Despite the lack of sequence similarity between MtmB and the methyltetrahydrofolate-binding domain of MetH, their overall structures are similar. Of this group of enzymes, the most mechanistically characterized system is the Mta complex catalyzing methanol-coenzyme M methyl transfer. As mentioned above, MtaC shows sequence homology with other corrinoid proteins involved in cytoplasmic methyl transfers to coenzyme M, and with the cobalamin-binding module of MetH. The active-site histidine responsible for the base-off,His-on ligation of the 5-hydroxybenzimidazolylcobamide corrinoid of MtaC was shown to be His136, the histidine in the signature AspX-His-X-X-Gly sequence [22]. MtaC is isolated in a complex with MtaB, and the complex was shown to catalyze methylation of the corrinoid prosthetic group using methanol as the methyl donor [32]. Recently, an X-ray structure of the MtaBC complex has been determined [33]. Thus far, this is the only structure of a methyl transferase corrinoidbinding protein in complex with one of its substrate binding domains. MtaC is indeed structurally related to the cobalamin-binding module of MetH, and as in that structure it contains both a four helix bundle (the cap) and the Rossmann domain responsible for cobalamin binding. As in the structure of the C-terminal fragment of His759Gly MetH, the cap is displaced to allow juxtaposition of MtaB with the Rossmann domain. MtaB is composed of an a8b8 barrel with similarities to the substrate-binding domains of MetH, and a unique helical layer that is not seen in MetH. A zinc atom is located at the C-terminus of the barrel in a deep funnelshaped pocket (Figure 10); this zinc was previously shown to be required for activity [34]. In the MtmBC structure the barrel is positioned over the Rossmann domain so as to position the zinc above the cobalt of the corrinoid prosthetic group and to define the methanol binding site at the interface. The catalytic zinc ion is ligated by Glu164, Cys220, and Cys269, and although it exhibits approximately tetrahedral geometry, is apparently lacking a fourth ligand. The authors assume that methanol binds to the fourth site on the zinc through its hydroxyl group. Additional electron density, which has been modeled as a potassium ion, is located 3.1 A˚ away from the zinc, and this putative potassium ion is also ligated by Glu164 as well as by other oxygen ligands. The authors suggest that the methanol will actually bridge between the potassium ion and the zinc. His136, the cobalt of the corrinoid, the methyl group and the oxygen of methanol and the catalytic zinc all form a line, favoring an SN2 mechanism in Met. Ions Life Sci. 2009, 6, 53–114


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Figure 10. Proposed catalysis of methyl transfer from methanol to the corrinoid cofactor in the MtmBC complex as deduced from the X-ray structure of the complex. The electron density indicated by X has been modeled as a potassium ion. Reproduced from [33] with permission from the National Academy of Sciences, USA, copyright 2006.

which the Co(I) state of the cofactor nucleophilically attacks the methyl group of methanol with the departing hydroxyl group remaining ligated to the zinc. In agreement with this proposed mechanism, which would result in inversion of configuration of the methyl group, the overall stereochemistry of the methyl group transfer from methanol to coenzyme M, which would require two successive nucleophilic displacements, proceeds with retention of stereochemistry [35]. MtaA catalyzes methyl transfer from MtaC to coenzyme M and also from exogenous methylcobalamin to coenzyme M [36]. Like the other proteins that catalyze alkyl transfer to thiols [37], MtaA contains a catalytically essential zinc ion that serves as the binding site for coenzyme M [38]. EXAFS analysis indicated that the substrate-free MtbA zinc was ligated by one sulfur ligand and three oxygen or nitrogen ligands, and that binding of coenzyme M was associated with replacement of one of the oxygen/nitrogen ligands by the sulfur of coenzyme M [39]. The MtaA isozyme MtbA also contains a catalytically essential zinc ion that serves as the binding site for coenzyme M [40]. EXAFS analysis indicated that the substrate-free MtbA zinc was ligated by two sulfur ligands and two oxygen or nitrogen ligands, and that binding of coenzyme M was associated with replacement of one of the oxygen/nitrogen ligands by the sulfur of coenzyme M [40]. These two isozymes share only 40% sequence identity, and presumably one of the cysteine ligands of MtaA is not conserved in MtaB. Met. Ions Life Sci. 2009, 6, 53–114

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2.4. Membrane-Associated Energy-Conserving Corrinoid Methyltransferase N5-Methyltetrahydromethanopterin:coenzyme M methyltransferase is an integral membrane protein that catalyzes an energy-conserving step in methane formation from CO2 and/or acetate in Methanobacterium thermoautotrophicum. The enzyme catalyzes methyl transfer from methyltetrahydromethanopterin, a methyl donor similar in structure to methyltetrahydrofolate, to the thiol of 0 coenzyme M. This reaction is exergonic (DG0 ¼ 30 kJ/mol) and is coupled to extrusion of a sodium ion across the cell membrane [41]. The enzyme was purified to homogeneity [42] and shown to comprise eight different subunits [43]. The corrinoid-containing subunit MtrA was intially identified as part of the complex by immunoprecipitation [44], and subsequently was shown to contain one equivalent of 50 -hydroxybenz-imidazolyl-cobalamide [45] bound with a histidine ligand to the cobalt [46]. The histidine ligand was shown to be His84 by mutagenesis, but interestingly there is no sequence homology between MtrA and the cobalamin-binding region of methionine synthase or the corrinoid proteins of the non-energy conserving cytoplasmic methyltransferases discussed in the previous section [47]. Mechanistic studies on this very large membrane-bound protein complex are challenging indeed. The Thauer laboratory succeeded in demonstrating that enzyme in the Co(II) form was inactive, and could be activated by incubation with titanium citrate and methyltetrahydromethanopterin, suggesting that the Co(I) form of the cofactor was required for reaction. Enzyme demethylation in the presence of coenzyme M did not require the presence of titanium citrate. The authors concluded that the basic mechanism was similar to that of MetH, in which the prosthetic group cycles in catalysis between cob(I)alamin and methylcobalamin forms [48]. As is the case with MetH and with the non-energy conserving cytoplasmic corrinoid methyltransferase, the complex can catalyze the methylation and demethylation of exogenous cobalamin as well as of the endogenous MtrA corrinoid. Only the half reaction involving transfer of the methyl group from the methylcobalamin to coenzyme M was sodium-ion dependent [49].

2.5. The Corrinoid Iron-Sulfur Protein Acetogenic bacteria like members of the genus Moorella produce acetate as the sole fermentation product during growth on either glucose or H2 and CO2. These organisms employ the Wood-Ljungdahl pathway (Figure 2), in which CO2 derived from acetate is converted to formate and then sequentially reduced to a methyl group bound to tetrahydrofolate in the ‘‘Eastern



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branch’’ of the pathway, and CO2 is converted to CO in the reaction catalyzed by CO dehydrogenase in the ‘‘Western branch’’ of the pathway. The corrinoid iron-sulfur protein accepts the methyl group of methyltetrahydofolate and transfers it to a nickel center on acyl-CoA synthase, which then catalyzes the carbonylation of the methyl-nickel bond and the subsequent transfer of the acetyl group to the thiol of coenzyme A [3]. The corrinoid iron-sulfur protein consists of two subunits. The large subunit, AcsC, contains a 4Fe-4S cluster, while both subunits are necessary for tight binding of the corrinoid prosthetic group. A separate methyltransferase, AcsE, catalyzes the methylation of the corrinoid using methyltetrahydrofolate as the methyl donor. Hydrogenogenic bacteria, which can utilize CO as a sole source of carbon and energy under anaerobic growth conditions, employ the Wood-Ljungdahl pathway for assimilation of carbon. They also contain a corrinoid iron-sulfur protein consisting of two subunits and a separate methyltransferase [50]. Methanogens also synthesize acetyl-CoA from methyltetrahydromethanopterin or methyltetrahydrosarcinopterin and CO2, but during methanogenesis they run the reaction in reverse, transferring the methyl group of acetyl-CoA to analogues of tetrahydrolate and generating carbon monoxide. The anabolic and catabolic acyl-CoA synthase/decarbonylase complexes appear to be similar, and a corrinoid has been shown to be involved in catabolism by the anabolic complex by inhibition with propyl iodide and relief of inhibition by photolysis [51,52]. Methanogens express a corrinoid iron-sulfur protein as part of this acetyl CoA decarbonylase/synthase multienzyme complex. As the name suggests, this complex catalyzes both acetyl-CoA synthesis and cleavage. Members of the genus Methanosarcina can grow on acetate by metabolizing it to CO2 and methane, with acetyl-CoA as an intermediate. The overall decomposition of acetyl-CoA is shown by equation (2), where H4SPT is tetrahydrosarcinapterin, an analogue of tetrahydrofolate, and H: represents CH3 CO-SCoA þ H4 SPT þ H2 O Ð CH3 -H4 SPT þ CO2 þ CoA þ H: ð2Þ hydride. The reducing equivalents produced in this reaction are used to reduce the methyl group of methyltetrahydrosarcinapterin to methane following transfer of the methyl group to coenzyme M [53]. Corrinoid iron-sulfur proteins have been isolated and characterized from Clostridium thermoaceticum (renamed Moorella thermoacetica) [1,54], from the methanogenic archaeon Methanosarcina thermophila [55] and from the hydrogenogenic bacterium Carboxydothermus hydrogenoformans [50]. The acetyl-CoA decarbonylase/synthase complex from Methanosarcina thermophila has been purified to homogeneity [56–58] and shown to contain the two subunits of the corrinoid iron-sulfur protein CdhE (the g subunit, 63 kDa) and


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CdhD (the d subunit, 53 kDa). These subunits show sequence homology to the two subunits, AcsC and AcsD, of the corrinoid iron-sulfur protein in acetogens [59] and to the CfsA and CfsB subunits from C. hydrogenoformans [60], although the smaller d subunit is considerably larger than the small subunits in acetogens and hydrogenogenic bacteria. The gd complex exhibits methyltransferase activity. In acetogens and hydrogenogenic bacteria, transfer of the methyl group from methyltetrahydrofolate to the corrinoid iron-sulfur protein requires a separate protein, but in methanogens this activity appears to be integrated into CdhD and/or CdhE. The spectroscopic properties of the corrinoid iron-sulfur protein from M. thermoacetica have been particularly extensively characterized. Tight binding of the corrinoid, which is 5-hydroxybenzimidazolyl-cobamide in M. thermoaceticum, requires the presence of both large and small subunits. The sequences of the genes specifying the two subunits have no homology with that specifying the cobalamin binding module of MetH and lack the DxHxxG motif associated with binding of cobalamin to MetH in the baseoff,His-on form [54]. The corrinoid is bound to the protein in the base-off form and its spectral properties indicate the absence of a nitrogen ligand in the lower axial position in both the methylcob(III)amide and cob(II)amide redox states of the corrinoid. In marked contrast, Maupin-Furlow and Ferry noted that the purified CdhD subunit of the M. thermophila protein could be reconstituted with hydroxocobalamin in the base-on configuration, while reconstitution of purified CdhE protein (the large subunit) led to base-off cobalamin binding [55]. The spectrum of the methylated protein was determined in the presence of sodium mersalyl, an organic mercurial reagent which disrupts the iron-sulfur center and allows unobstructed study of the cobalt chromophore. The spectrum exhibits a peak at B440 nm, which is consistent with a base-off methylcorrinoid, and release of the cofactor from the protein leads to an absorbance spectrum with a maximum of 537 nm, typical of a base-on methylcorrinoid [1]. The enzyme is isolated in the cob(II)amide form, and its EPR spectrum exhibits eight singlet peaks centered around g ¼ 2 [1,54]. This spectrum is diagnostic of base-off cob(II)amides, and is due to the hyperfine splitting imposed by the spin 7/2 of the cobalt nucleus. The absence of superhyperfine splitting, which would lead to an octet of triplets rather than singlets, demonstrates that nitrogen is not coordinated to 5-hydroxybenzimidazolylcobamide in the protein. Similar results have been observed for the corrinoid/iron-sulfur component of the acyl-CoA synthase/decarbonylase complex from Methanosarcina thermophila [61]. In a collaboration between the laboratories of Thomas Brunold and Stephen Ragsdale, the axial ligation of the corrinoid cofactor has been determined using a combination of magnetic circular dichroism, EPR, Met. Ions Life Sci. 2009, 6, 53–114


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resonance Raman and computational chemistry [62]. This analysis demonstrated that both the methylcob(III)inamide and cob(II)inamide states of the prosthetic group are present in the base-off form, with an axial water ligand. These spectroscopies cannot distinguish between b and a axial ligation of the water in the cob(II)inamide state, but presumably the water is in the a position when the prosthetic group is methylated. The role of the iron-sulfur center of the corrinoid iron-sulfur protein was also examined in the M. thermoacetica enzyme. The cobalt(I) form of the enzyme undergoes oxidation to form an inactive cobalt(II) species about once in every 100 turnovers [63]. The iron-sulfur center of the corrinoid ironsulfur protein has been shown to be required for reductive reactivation of the inactivated cob(II)amide prosthetic group, but is not required for the methyl transfers catalyzed by the active enzyme [64]. This cluster has a redox potential of 523 mV versus the standard hydrogen electrode [59], which is nearly isopotential with the cob(II)amide/cob(I)amide couple of the enzymebound corrinoid. For this reason, efficient reduction does not require coupling to ATP hydrolysis or methyl transfer from AdoMet. The oxidized ironsulfur cluster can then be reduced by carbon monoxide/carbon monoxide dehydrogenase, by hydrogen/hydrogenase, or by a low-potential reduced ferredoxin [64]. The structure of the corrinoid iron-sulfur protein from C. hydrogenoformans was recently determined [60]. This structure revealed that the large subunit has three domains: an N-terminal domain that binds the 4Fe-4S cluster, a central a8b8 barrel, and a C-terminal domain that binds the cobalamin in the expected ‘‘base-off’’ conformation. The small subunit was also an a8b8 barrel. The small subunit packs against the upper face of the cobalamin prosthetic group. This binding mode for the cobalamin is in agreement with earlier observations that the M. thermoacetica corrinoid is bound most tightly when both subunits are present [54]. No protein ligand to the cobalamin was identified, consistent with its binding in a base-off conformation, but a water ligand was seen in the upper axial (b) position. The structure could not distinguish between cobalt in the +2 or +3 oxidation state of the cofactor. The prosthetic group ligation seen in this structure is in excellent agreement with the spectroscopy reported for the M. thermoacetica enzyme [62], and with the previously puzzling observations of MaupinFurlow and Ferry on the M. thermophila protein, who observed base-on binding of hydroxocobalamin to the large subunit and base-on binding to the small subunit [55]. In a DALI search (a program that looks for structural similarity in proteins) to identify structures with high similarity to the subunits of the corrinoid iron-sulfur protein, the highest similarity was found to be with the folate-binding domain of MetH, which is similar to both of the a8b8 barrels. Structural similarity was also seen between these two barrels and the Met. Ions Life Sci. 2009, 6, 53–114

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methyltransferase AcsE from M. thermoacetica [65]. The C-terminal domain of the large subunit was also found to exhibit structural similarity to the cobalamin-binding domain of MetH. Catalysis of the methyl transfers from methyltetrahydrofolate or its analogues to the nickel site of acyl-CoA synthase requires that the corrinoid iron-sulfur protein interact first with the methyltransferase to receive the methyl group of methyltetrahydrofolate and then with the metal site on acylCoA synthase to form methyl-Ni. Reactivation of the inactive cobalt(II) form of the enzyme presumably requires a third conformation in which the iron-sulfur center is juxtaposed with the b-face of the corrinoid prosthetic group. Thus the corrinoid iron-sulfur protein must undergo a series of conformational changes that may resemble those seen for MetH. What advantages does the base-off state of the prosthetic group confer to the corrinoid iron-sulfur protein? Base-off cob(II)alamin, formed at low pH, is more facilely reduced than base-on cob(II)alamin [66] and the enzymebound 5-methoxybenzimidazolylcob(II)amide is also more readily reduced than the free base-on cofactor [59], as is the enzyme-bound cob(II)alamin found in the M. thermophila [61] and M. barkeri [57] proteins. In the case of methionine synthase, the cob(II)alamin form of the enzyme can be readily interconverted between base-on and base-off forms, while the methylcobalamin form of the enzyme is present predominantly as the base-on form. However, the methylcobinamide form of the corrinoid iron-sulfur protein is completely base-off. The acetogenic acyl-CoA synthase shares another property with methionine synthase and other corrinoid-dependent methyltransferases, namely the ability to react with exogenous corrinoids as well as with their physiological corrinoid partner proteins. The Ragsdale group has shown that acyl-CoA synthase from M. thermoacetica reacts 2000-times faster with methylcobinamide, which lacks a lower axial base, than with methylcobalamin [67]. These results may suggest a late transition state for this methyl transfer, with substantial bond breaking leading to cob(I)inamide character.

2.6. Aromatic O-Demethylases Many acetogenic bacteria derive both carbon and energy by demethylating aromatic methyl ethers. The pathway involves transfer of the methyl group from the ether to methyltetrahydrofolate, which then can enter the Western branch of the Wood-Ljungdahl pathway and be converted into the methyl group of acetate. Methyl groups bound to methyltetrahydrofolate can also be oxidized to formate and/or CO2 by the reverse of the Eastern branch of the Wood-Ljungdahl pathway. The reversal of the Eastern branch to CO2 generates three hydride equivalents per methyl group oxidized, which in turn Met. Ions Life Sci. 2009, 6, 53–114


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can be used to reduce 3 moles of CO2 to CO to form 3 mol of acetyl-CoA in the Western branch. The overall stoichiometry is shown in equation (3). 4ROCH3 þ 2CO2 þ 3CoASH ! 4ROH þ 3CoAS-CðOÞ-CH3 þ H2 O ð3Þ The O-demethylases from Acetobacterium dehalogenans [68,69] and Moorella thermoacetica [70] conform to the basic pattern illustrated in Figure 3. The four components of the vanillate O-demethylase from A. dehalogenans comprise two methyltransferases, a corrinoid protein and an activating enzyme [68,69]. The first methyltransferase (odmB) catalyzes methyl transfer from vanillate (O-methylhydroxybenzoate) to the corrinoid protein (odmA) and the second methyltransferase catalyzes methyl transfer from the methylated corrinoid protein to tetrahydrofolate. The activating protein uses hydrogenase as the source of reducing equivalents and requires ATP. The amino acid sequence derived from the odmA gene shows about 60% similarity with the cobalamin-binding domain of methionine synthase, including the Asp-x-His-x-x-Gly sequence characteristic of binding cobalamin in the base-off,His-on mode.

2.7. Reductive Dehalogenases Free cob(I)alamin is known to react rapidly with alkyl halides like methyl iodide to produce alkyl cobalamins, and in the presence of reducing agents abiotic dehalogenation is catalyzed at significant rates [71]. The mechanism of abiotic reductive dechlorination of perchloroethylene has recently been examined [72]. The authors concluded that the most likely mechanism was that shown in Figure 11. This mechanism involves an initial electron transfer from cob(I)alamin to the perchloroethylene with release of chloride ion and formation of a trichlorovinyl radical that would immediately combine with cob(II)alamin to produce a trichlorovinylcobalamin. A further electron transfer would then generate a trichlorovinyl anion and regenerate cob(II)alamin.

Figure 11. Proposed abiotic mechanism for dechlorination of perchloroethylene. Redrawn from [72] with permission from the American Chemical Society, copyright 2005. Met. Ions Life Sci. 2009, 6, 53–114

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With this mechanism as a guide, we can now examine the studies on corrinoid-dependent enzymes that catalyze dehalogenation reactions in anaerobic bacteria. Many of the organisms capable of aryl halide conversion belong to the sulfidogenic bacteria, which can reductively dechlorinate polychlorinated phenols and benzoates, tetrachloroethylene and trichlorethylene (reviewed in [73]). The perchloroethylene reductive dehalogenases have been particularly well characterized. These enzymes are typically membrane anchored, and contain two Fe4S4 or Fe3S4 clusters in addition to a corrinoid, which is present in the base-off form [74,75]. The redox potentials of the iron sulfur centers are lower than that of the Co(II)/ Co(I) couple of the corrinoid, which would favor reduction of the corrinoid [74,75]. An ortho-chlorophenol reductive dehalogenase was purified from Desulfitobacterium dehalogenans and shown to have similar properties [76], including the presence of base-off corrinoid as isolated in the Co21 oxidation state. This protein contained one Fe4S4 and one Fe3S4 cluster, with the Fe4S4 cluster having the lower potential. More recently a meta- and para-chlorophenol reductive dehalogenase was purified, cloned and sequenced from Desulfitobacterium frappieri [77]. The CprA protein sequence contains two iron-sulfur binding motifs, and a Glu-Tyr-His-Tyr-Asn-Gly motif (EYHYNG) that is related to the Asp-x-His-x-x-Gly motif found in base-off, His-on cobalamin-binding proteins. It will be of great interest to know whether this histidine is indeed a ligand to the cobalt under some conditions. Given the reaction mechanism proposed in Figure 11, binding of cobalamin in the base-off mode should greatly facilitate the reduction of both cob(II)alamin and trichlorovinyl-cobalamin. The enzymes responsible for reductive dehalogenation of chloromethane have also been studied. These enzymes catalyze methyl transfer from chloromethane to tetrahydrofolate. In Methylobacterium sp., the reductive dehalogenase comprises two components, CmuA and CmuB. CmuA is a two domain methyltransferase/corrinoid-binding protein and contains cob(II)alamin as isolated [78]. The N-terminal domain shows sequence similarity to methycobamide:coenzyme M methyltransferases, while the C-terminal domain shows sequence similarity to the cobalamin-binding domain of methionine synthase. However the Asp-x-His-x-x-Gly sequence characteristic for base-on,His-off cobalamin binding is replaced by Asn-Thr-Gln-x-x-Gly in the sequence alignment, suggesting a base-off mode of cobalamin binding [79]. CmuB shows methylcobalamin:tetrahydrofolate methyltransferase activity and shares sequence similarity with MtrH of the methyltetrahydromethanopterin:coenzyme M methyl transferase complex and with the methyltetrahydrofolate-binding domain of MetH [80]. There is no evidence for iron-sulfur cluster motifs in the sequences of CmuA or CmuB and the spectra of the enzymes as isolated, or after reduction, do not reveal the presence of iron-sulfur clusters. Met. Ions Life Sci. 2009, 6, 53–114


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2.8. Modes of Activation of Corrinoid-Dependent Methyltransferases A majority of the corrinoid-dependent methyltransferases requires a reductive activation, even those proteins native to strictly anaerobic organisms. In MetH, reductive activation is necessary to retrieve the inactive cob(II)alamin prosthetic group and return it to the catalytic cycle, but it is less certain whether such oxidative inactivation actually occurs under strictly anaerobic growth conditions. Recent studies on the incorporation of the cobalamin cofactor into the apoenzyme suggest an alternative function for these activation proteins or modules, namely a role as a chaperone to assist the enzyme in binding the prosthetic group [81–83]. In the methyltransferases that bind the cobalamin prosthetic group in the base-off or base-off,His-on state, incorporation of the prosthetic group requires that the dimethylbenzimidazole first be dissociated from the cobalamin. The free energy required for replacement of dimethylbenzimidazole by water has been measured for a series of cobalamins [84,85]. For aquacob(III)alamin, the base-off form is disfavored by 10.4 kcal/mol, while for cob(II)alamin, the base-off form is only disfavored by 3.3 kcal/mol. Thus on thermodynamic grounds alone, we would expect the initial formation of base-off cobalamin to occur at the cob(II)alamin oxidation state and to be an endergonic process. Table 1 summarizes what is known about the activation systems in corrinoid-dependent methyltransferases. I will again begin by discussing what is known about the activation of methionine synthase, and then proceed to describe studies on the ATP-dependent activation systems found in other corrinoid-dependent methyltransferases. The activation of the corrinoid iron-sulfur protein, which does not require either ATP or AdoMet, has already been discussed in Section 2.5. In MetH from E. coli, activation of enzyme in the cob(II)alamin form requires AdoMet [86] and a reducing system. While early studies of the enzyme employed exogenous reductants, isolation of the components responsible for reduction in crude bacterial extracts in the laboratory of Frank Huennekens led to the identification of flavodoxin, flavodoxin (ferredoxin) reductase, and NADPH as the components responsible for MetH reduction in these extracts [87]. Incubation of MetH, isolated in the cob(II)alamin form with AdoMet bound, with flavodoxin, flavodoxin reductase and excess NADPH led to formation of enzyme in the methylcobalamin form [88]. As mentioned previously, AdoMet is bound to the C-terminal module of methionine synthase [6,14], which is required for reductive activation. This C-terminal module also contains determinants for the binding of flavodoxin [89]. In 1997, Hoover et al. reported that incubation of MetH in the cob(II)alamin form with oxidized flavodoxin led to the formation of base-off Met. Ions Life Sci. 2009, 6, 53–114

ATP

ATP/reduced ferredoxin

ATP H2/hydrogenase ferredoxin CO/CO dehydrogenase ferredoxin

MAP

RamA (MA0150)

not identified

not identified

requires Fe4S4 center on large subunit

Methanosarcina barkeri

Methanosarcina barkeri Methanosarcina acetovorans Methanobacterium thermoautotrophicum

Acetobacterium dehalogenans Moorella thermoacetica

Methanol:coenzyme M methyltransferase (5-OHbenzimidazolylcobamide) Methylamine:coenzyme M methyltransferases (5-OHbenzimidazolylcobamide) Energy-conserving methyltetrahydromethanopterin: coenzyme M methyltransferase (5-OH-benzimidazolylcobamide) Veratrol:H4folate and vanillate:H4folate Omethyltransferases (cobalamin) Corrinoid Fe-S protein (5methoxybenzimidazolyl)

AdoMet/flavodoxin AdoMet/methionine synthase reductase ATP/H2/hydrogenase

Cofactors/Reductants

AdoMet domain AdoMet domain

Escherichia coli Homo sapiens

Methionine synthase (cobalamin)

Organism

Activating Protein(s) or Domains

Requirements for reductive activation of corrinoid-dependent methyltransferases.

Enzyme Complex (Corrinoid Cofactor)

Table 1.

Met. Ions Life Sci. 2009, 6, 53–114 [63]

[95,96]

[94]

[28]

[83,93]

[87] [90,92]

References

80 MATTHEWS


81

cob(II)alamin [9]. This shift required stoichiometric concentrations of methionine synthase and flavodoxin. When the structure of the C-terminal half of His759Gly methionine synthase in the AdoMet:Cob conformation was determined, it was apparent that assumption of this conformation led the cobalamin to assume a base-off conformation. Thus oxidized flavodoxin, which is incapable of electron transfer, is assuming a role as a chaperone to facilitate the formation of base-off cobalamin in the AdoMet:Cob conformation when the enzyme is in the cob(II)alamin form. This is of course the conformation needed for reductive remethylation of the cofactor. If the enzyme is in the aquacob(III)alamin form, flavodoxin binds tightly, but the shift to the base-off conformation does not occur. Mammals lack flavodoxin and flavodoxin (ferredoxin) reductase. For the activation of methionine synthase they instead employ a fusion protein with an N-terminal domain homologous to flavodoxin and a C-terminal domain homologous to flavodoxin reductase [90]. This protein, methionine synthase reductase, is required for the in vivo activation of methionine synthase, and patients with severe deficiencies of this enzyme present with symptoms resembling those of patients with severe deficiencies of methionine synthase itself [91]. Methionine synthase reductase is not only required for the reductive reactivation of human methionine synthase. The oxidized reductase substantially stabilizes the apoenzyme, which rapidly undergoes irreversible denaturation on incubation at 37 1C [82]. Furthermore, methionine synthase reductase greatly increases the yield of holoenzyme formed on incubation of apo-methionine synthase with aquacobalamin and dithiothreitol [82]. These findings suggest a chaperone-like function for methionine synthase reductase. Thus far, insufficient amounts of human methionine synthase have been available to permit experiments to determine whether methionine synthase reductase also stabilizes holoenzyme in a base-off AdoMet:Cob conformation. Thus far, AdoMet-dependent reductive activation appears to be unique to methionine synthase. Where activation of other methyltransferases has been shown to occur, reductive activation appears to be coupled to ATP rather than AdoMet. For a listing of methyltransferase activating proteins see Table 1. The best-studied reactivation protein is the methyltransferase-activating protein (MAP) which is involved in the activation of the cytoplasmic methylamine:coenzyme M methyltransferases and methanol:coenzyme M methyltransferase in Methanosarcina barkeri. Sustained activity of these coenzyme M methyltransferases requires ATP, hydrogen, hydrogenase and MAP [83]. Incubation of ATP with MAP at 1:1 concentration ratios led to the phosphorylation of MAP. Phosphorylated MAP substituted for ATP in the stoichiometric activation of MT1, the corrinoid-containing methanol:5hydroxybenzimidazolyl cobamide component of the cytoplasmic methanol:coenzyme M methyltransferase complex. If ATP were present in excess, Met. Ions Life Sci. 2009, 6, 53–114

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MAP could catalyze multiple rounds of activation, indicating that ATP hydrolysis occurred during the activation process. If MT1 in the aquacob(III)inamide form was incubated with hydrogen and hydrogenase, base-on cob(II)amide was formed. Addition of MAP and ATP resulted in formation of a mixture of base-on (60%) and base-off (40%) cob(II)inamide, and if methanol was then added, the prosthetic group was quantitatively converted to the methylcobinamide form [93]. Thus, phosphorylated MAP acts as a chaperone, inducing a conformation change in MT1 that favors the formation of base-off cob(II)inamide, similar to the effect of oxidized flavodoxin on MetH in the cob(II)alamin form. The difference is that methanol, the substrate, also serves as the methyl donor in reductive reactivation. A protein thought to be similar or identical to MAP can also serve to activate the corrinoid-proteins in the methylamine methyltransferase complexes [97]. However, annotation of the Methanosarcina acetivorans genome sequence [28,98] indicates that a gene specifying an iron-sulfur protein designated RamM is responsible for activation of methylamine methyltransferases. A number of homologues of ramM have been identified in the M. acetivorans genome, although their functions have not yet been determined.

2.9. Methyl Transfer in Fosfomycin Biosynthesis The biosynthesis of fosfomycin was initially proposed to involve a unique function for methylcobalamin, namely the transfer of the methyl group as a methyl anion (Figure 12). However, all characterized methylcobalamin-dependent methyltransferases transfer the methyl group as a methyl cation. The

Figure 12. Originally proposed mechanism for generation of the methyl group of fosfomycin. Adapted from [101] with permission from the Royal Society of Chemistry, copyright 2007. Met. Ions Life Sci. 2009, 6, 53–114


83

evidence for the involvement of methylcobalamin as the methyl donor comes from the studies of Seto and Kuzuyama and their colleagues [99,100]. They showed that a mutant strain of Streptomyces with a block in the B12 biosynthetic pathway could not produce fosfomycin, and that feeding of 14C-labeled methylcobalamin to this blocked mutant resulted in 14C-labeled fosfomycin. Recently, van der Donk and his colleagues [101] identified the entire biosynthetic cluster for fosfomycin and showed that it induced fosfomycin production in a Streptomyces strain lacking this capability. Analysis of the gene sequence of one of the components of the cluster, fom3, showed that it contained a region with similarities to a B12-binding domain and another region with homology to the family of proteins that utilize AdoMet as a radical generator. The fom3 gene was shown to be essential for the biosynthesis of fosfomycin. Furthermore the authors provided a strong inference that the actual substrate for methylation was hydroxyethylphosphonate rather than phosphonoacetaldehyde, leading to the proposal of a mechanism much more in keeping with known B12 chemistry (Figure 13). We must await the purification and characterization of Fom3. But if the proposed mechanism is indeed correct, we may have a fascinating clue to the origin of AdoCbl-dependent enzymes. As pointed out by Sauer and Thauer in their review [53], thus far all corrinoid protein characterized from methanogenic Archaea have been methyltransferases containing methylcobalamin as a prosthetic group and no AdoCbl-dependent enzymes have been found. Furthermore, the cobO gene required for synthesis of AdoCbl appears to be lacking. Fom3, and its analogues in methylation reactions required for the biosynthesis of other antibiotics produced in Streptomyces, may represent a

Figure 13. Mechanism for generation of the methyl group of fosfomycin proposed by van der Donk and colleagues. Adapted from [101] with permission from the Royal Society of Chemistry, copyright 2007. Met. Ions Life Sci. 2009, 6, 53–114

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step in the direction of development of the AdoCbl family of enzymes. In the mechanism proposed for Fom3, AdoMet is cleaved to generate an adenosyl radical which abstracts a hydrogen from hydroxyethylphosphonate. The substrate radical is then methylated by methylcobalamin leaving cob(II)alamin as the product. Regeneration of methylcobalamin might then occur by way of a process similar to the reactivation of methionine synthase, using an external reductant and AdoMet as the methyl donor. It should be noted that this proposed mechanism would require two molecules of AdoMet per methylation reaction: one to generate the initial radical and a second to serve as a methyl donor.

3. ADENOSYLCOBALAMIN-DEPENDENT REARRANGEMENTS AND ELIMINATIONS Although the AdoCbl-dependent enzymes initially attracted the greatest attention from organic and inorganic chemists because of their fascinating chemistry, we now know that they represent but a small branch of the corrinoid-dependent enzymes. They are found most frequently in eubacteria (Domain Prokaryota) and just one AdoCbl-dependent enzyme, methylmalonyl-CoA mutase, is found in mammals. No AdoCbl-dependent enzymes have yet been identified in Archaea. The basic mechanism of AdoCbl-dependent rearrangements is shown in Figure 14. This mechanism was simultaneously elucidated in Abeles’ laboratory at Brandeis University and Arigoni’s laboratory in Zu¨rich. In classic papers, Frey and Abeles [102] showed that AdoCbl bound to propanediol dehydrase is tritiated as the enzyme reacts with [1-3H]1,2-propanediol and that tritium could subsequently be transferred from the isolated tritiated coenzyme to unlabeled propanediol, and Re´tey and Arigoni [103] then showed that AdoCbl that had been labeled with tritium when catalyzing the propanediol dehydrase reaction could also transfer tritium to methylmalonylCoA. Frey, Essenberg, and Abeles [104] showed that tritium is transferred from [1-3H]propanediol to the C-5 0 position of the AdoCbl of diol dehydrase, and from [5 0 -3H]AdoCbl to C2 of the product propionaldehyde. They also showed that the hydrogen abstracted from C1 of the substrate becomes equivalent with one or both of the hydrogens of the C5 0 position of the cofactor following the initial hydrogen transfer. Cleavage of the Co-C5 0 bond of AdoCbl to form cob(II)alamin and 5 0 -deoxyadenosine was first observed in the suicide inactivation of diol dehydrase by glycolaldehyde, and in the same paper the substrate propanediol was also shown to induce transient formation of cob(II)alamin [105]. At the same time, Re´tey, Umani-Ronchi, Seibl, and Arigoni [106], used 18O-enriched



Figure 14.

85

Basic mechanism of AdoCbl-dependent rearrangements.

propanediols to demonstrate the migration of 18O from C2 of propanediol to C1 of propionaldehyde, thus demonstrating the migration of the substituent at C1 (X in Figure 14) in another classic paper. In the sections that follow, I will review the literature on each of the AdoCbl-dependent enzymes in turn. First, however, I wish to emphasize several challenges to understanding the mechanisms of each of these enzymes: Activation of the C-Co bond of AdoCbl for cleavage: The carboncobalt bond of AdoCbl is estimated to have a bond dissociation energy of 30 kcal/mol [107]. Thus Finke and Hay have estimated that diol dehydrase must lower the barrier for Co-C bond homolysis by at least 14.7 kcal/mol for a rate acceleration of 1010! How this is achieved in any AdoCbl-dependent enzyme remains controversial. Transfer of H from substrate to deoxyAdo and from deoxyAdo to product: For some of the AdoCbl-dependent enzymes, the H must traverse long distances (6–10 A˚) during the reaction. We are just now beginning to understand how H is transferred by the various enzymes. Catalysis of the migration of X: The issue of whether cob(II)alamin is a participant in catalysis or a bystander is an argument that has persisted over decades. However, recent studies have greatly illuminated the mechanisms of migration, especially in the enzymes that catalyze carbon skeleton rearrangements. It has become clear that the distance between cob(II)alamin and the substrate radical following homolytic cleavage of AdoClb in diol dehydrase and ethanolamine ammonia lyase is too great for the cobalamin to participate in the subsequent rearrangment. However, in the mutases, the distance between cob(II)alamin and


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substrate radical would be consistent with participation of the cobalamin in the rearrangement [108], and in fact density functional theory calculations support this role for cobalamin [109]. In reviewing the voluminous literature on these enzymes, I have tried to emphasize recent developments in the field, particularly emphasizing what we have learned as X-ray structures of these proteins have been determined. But I have not done justice to the earlier stereochemical experiments that elucidated the details of the overall reactions, nor the extensive characterization of the substrate and product radicals. Rather, I have attempted to focus on the role of the B12 cofactor.

3.1. Enzymes Catalyzing Carbon Skeleton Rearrangements This class of enzymes catalyzes rearrangements that require cleavage of a carbon-carbon bond to allow migration of X. These enzymes are glutamate mutase, methylmalonyl-CoA mutase, and isobutyryl CoA mutase. In all three of these enzymes, the AdoCbl cofactor is ligated by a histidine residue from the protein. Indeed, following the cloning of glutamate mutase, Marsh and Holloway first recognized the Asp-X-His-X-X-Gly motif that characterizes His-on ligation in methionine synthase and the enzymes that catalyze carbon skeleton rearrangements [12]. While all three enzymes have similar cobalamin-binding domains or subunits, they differ considerably in the structures/sequences of the substrate-binding regions of the proteins.

3.1.1. Glutamate Mutase Glutamate mutase catalyzes the reaction shown in equation (4), the conversion of (S)-glutamate to (2S,3S)-3-methylaspartate. In this reaction, –

H3N COO H – OOC Cβ Cα H H H H

+

+

H3N

–

OOC

COO– H Cβ H

H Cα

H

ð4Þ

H

a hydrogen on Cb is exchanged with the glycyl group on Ca. The enzyme is an a2b2 oligomer; the small subunit GlmS (s) is 14.7 kDa while the large



87

subunit GlmE (e) is 50 kDa [110]. As noted above, the small subunit contains the Asp-X-His-X-X-Gly motif associated with binding of B12 in a DMB-off,His-on state. However, neither the large nor the small subunit binds AdoCbl tightly, and the binding site is created at the interface between the two subunits. For some studies, a fusion protein comprising both large and small subunits has been used to avoid the complications of subunit dissociation and loss of activity and AdoCbl [111]. The structure of glutamate synthase with AdoCbl substituted by methyl- or cyanocobalamin reveals that the architecture of GlmS is highly similar to that of the B12-binding domain of methionine synthase from E. coli [112]. The DMB nucleotide is deeply buried in a hydrophobic pocket in this subunit, and His16, which is carried on the loop between the first strand and the first helix of the barrel, coordinates the a position of the cobalt in B12. As suggested by the conserved motif, His16 is also hydrogen bonded to Asp14; however, the third amino acid in the ‘‘ligand triad’’ is missing and instead Asp14 also forms hydrogen bonds to main-chain amide groups and to a water molecule. A structure of GlmS apoenzyme has been determined by NMR and provides insights into how the holoenzyme may be formed [113]. In this structure, which otherwise resembles the architecture of the holoenzyme small subunit, residues 13–27 form a disordered and highly mobile loop. The region corresponding to the first a-helix in the holoenzyme, residues 18–27, rapidly interconverts between unstructured forms and an a-helical conformation. The unfolding of the first helix exposes to solvent the cavity where the DMB will reside in the holoenzyme, so that the apoenzyme is preorganized for incorporation of the B12 cofactor. I have long argued that only cobalamins with relatively high propensities to form base-off cobalamin (e.g., methyl- and adenosyl-cobalamin, cob(II)alamin, and cob(I)alamin) will lend themselves to incorporation into proteins that bind the cofactor in the base-off,His-on state, but this NMR structure adds another element to our understanding of the process by which holoenzyme formation might occur. The GlmE subunit is an a8b8 barrel, with the open end packed against the b face of the B12 in the holoenzyme structure [112]. The crystallization medium contained tartrate, an analogue of methylaspartate, which bound in the barrel in close proximity to the B12 cofactor. In a subsequent paper [114], the structure of active glutamate mutase with AdoCbl and glutamate bound was determined. In this structure, the electron density of the adenine is clearly modeled, but fitting the electron density with the ribose moiety of adenosine requires modeling in a mixture of C2 0 -endo and C3 0 -endo conformations (Figure 15). The C3 0 -endo conformation places C5 0 of the ribose within bonding distance of the cobalt of cobalamin, while the C2’-endo conformation leads to a 4.2 A˚ distance between C5 0 and Co, but places C5 0


88

MATTHEWS

Figure 15. Alternate conformations of the ribose of AdoCbl at the active site of glutamate mutase. On the left the ribose assumes a C3 0 -endo conformation, placing the 5 0 -carbon within bonding distance of the cobalt of cobalamin and distant from Cg of the glutamate substrate. On the right the ribose assumes a C2 0 -endo conformation, leading to breakage of the carbon-cobalt bond and placing C5 0 within van der Waal’s distance of Cg of the substrate.

within 3.3 A˚ of Cg of the glutamate substrate. Thus the authors propose that a simple pseudorotation of the ribose between two low-energy conformers leads to cleavage of the carbon-cobalt bond and abstraction of hydrogen from Cg of glutamate. Figure 16 shows the mechanism proposed for glutamate mutase. The initial research supporting this mechanism was performed in Horace Barker’s laboratory in Berkeley and has been recently reviewed [110]. These studies established the stereochemistry of the reaction, and showed that there was no exchange of the hydrogens of substrate or product with solvent, and no exchange of potential intermediates such as glycine or acrylate. The reaction is unique among the enzymes catalyzing rearrangement of carbon skeletons in that the migrating carbon is sp3 hybridized, as shown in equation (4). Initial

Figure 16. Proposed mechanism for glutamate mutase. The adenosyl radical formed by cleavage of the C-Co bond of AdoCbl abstracts a hydrogen from glutamate to yield the glutamyl radical. This leads to the elimination of acrylate and formation of glycyl radical, which in turn condenses with acrylate to form the b-methylaspartate radical. In the final step, the product radical abstracts hydrogen from 5 0 -deoxyadenosine to regenerate the adenosyl radical, which can now recombine with cob(II)alamin to reform AdoCbl. Met. Ions Life Sci. 2009, 6, 53–114


89

evidence for the fragmentation to form a glycyl radical and acrylate came from the observation that glycine and acrylate inhibit the enzyme synergistically, and induce the formation of an EPR spectrum similar to those observed for the substrate or product radical (reviewed in [110]). More recently, direct evidence for formation of glycine and acrylate was obtained by rapid-quench analysis of the reaction, and the rate of formation of these intermediates was shown to be faster than the overall rate of reaction [115].

3.1.2. Methyleneglutarate Mutase Methyleneglutarate mutase catalyzes a reaction that is very similar to that catalyzed by glutamate mutase, as shown in equation (5). One key difference however, in the reaction is that the migrating carbon is sp2 hybridized, a

H

H

H

H –

OOC

COO–

–

H COO–

OOC H

methyleneglutarate

H

ð5Þ

H

(R)-3-methylitaconate

property that is shared with all the other enzymes that catalyze carbon skeletal rearrangement except glutamate mutase. This property permits the rearrangement to proceed through a cyclopropylcarbinyl intermediate rather than requiring a fragmentation-recombination as shown in Figure 17 [110]. Rearrangements of cyclopropinyl radicals are well precedented in model chemistry and proceed extremely rapidly [116].

Figure 17. Mechanism proposed for radical rearrangment in methyleneglutarate mutase. Because the migrating carbon is sp2, a cyclopropylcarbinyl radical can form to mediate the transfer of the radical between Ca and Cb [116]. Met. Ions Life Sci. 2009, 6, 53–114

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The protein is isolated from Clostridium barkeri as a homotetramer of 60 kDa subunits. The deduced amino acid sequence of the protein shows significant sequence homology in its C-terminal region with the cobalaminbinding regions of methylmalonyl-CoA mutase, glutamate mutase and MetH, including the conserved Asp-X-His-X-X-Gly sequence that is the hallmark of DMB-off,His-on binding of the cobalamin cofactor [117]. Mutation of the corresponding residues, His485 and Asp483, decreases the rate of substrate turnover by 44000-fold and by 2000-fold respectively [118].

3.1.3. Methylmalonyl-Coenzyme A Mutase Methylmalonyl-CoA mutase catalyzes the reaction shown in equation (6). The migrating carbon is sp2, allowing radical rearrangement to proceed

ð6Þ

by way of a cyclopropylcarbinyl radical, as in methyleneglutarate mutase. The bacterial enzymes are ab heterodimers, with considerable homology between the a and b subunits, while the mammalian enzyme is an a2 homodimer. Only one molecule of AdoCbl is bound per bacterial heterodimer, however. The X-ray structure of the enzyme from Propionibacterium shermanii was the first structure to be determined of a complete cobalaminbinding protein [119]. The a- and b-chains exhibit similar folds, but only the a-chain contains bound cofactor. Each chain consists of an N-terminal a8b8 barrel and a C-terminal domain that exhibits a fold similar to the cobalaminbinding domain of methionine synthase. Ne of HisA610 coordinates the lower axial position of the B12 cofactor, which appears to be cob(II)alamin in this structure. Nd1 of HisA610 is hydrogen bonded to the carboxyl oxygen of AspA608, and the other carboxyl oxygen of the Asp608 side chain is hydrogen bonded to LysA604. As in glutamate mutase and MetH, the DMB substituent of the corrin ring is deeply buried in a hydrophobic pocket in the cobalamin-binding domain. The N-terminal barrel of the a-chain of methylmalonyl-CoA mutase is juxtaposed against the b-face of the B12, similar to its position in glutamate mutase [119]. The protein was crystallized in the presence of desulfo-CoA, a



91

substrate analogue that lacks the terminal thiol and the succinyl group of succinyl-CoA. One equivalent of this analogue was bound in a narrow tunnel along the axis of the barrel of the a-chain, completely buried in the interior of the barrel [119]. Structures of methylmalonyl-CoA mutase were subsequently obtained for the substrate-free enzyme and for enzyme in a non-productive complex with CoA [120]. These structures, which were similar, revealed that in the absence of productively bound CoA, the a8b8 barrel is split apart and the CoA binding site is accessible to solvent. When CoA binds, the barrel closes up and encapsulates the substrate. The adenosyl group of AdoCbl could be seen in the substrate-free complex, but when the active site closes it is no longer visible, and the TyrA89 side chain now occupies a position that overlaps with the adenosyl binding region in the substrate-free enzyme. The authors propose that the closing of the active site cavity forces the carbon-cobalt bond cleavage. Support for the role of TyrA89 as a ‘‘molecular wedge’’ comes from studies of TyrA89Phe and TyrA89Ala mutant enzymes that demonstrate 1000-fold reductions in catalytic activity, and the disappearance of cob(II)alamin from the spectrum of the enzyme taken under steady-state conditions [121]. In the wild-type enzyme the ratio of AdoCbl:cob(II)alamin is about 4:1 when the enzyme is catalyzing the conversion of methylmalonyl- to succinyl-CoA. These studies enforce the view that the enzyme uses conformational changes driven by the binding of the CoA substrate to break the carbon-cobalt bond of the cofactor, consistent with the earlier observation that substrate binding accelerates carbon-cobalt bond cleavage by a factor of 1012 [122]. Padmakumar and Banerjee have measured the Co-C homolysis rate of AdoCbl bound to methylmalonyl-CoA mutase [122]. When the rates of homolysis are compared in the presence of [CH3]methylmalonyl-CoA and [CD3]methylmalonyl-CoA, the rate in the presence of the deuterated substrate is at least 20-fold slower. One would not expect the deuteration of the substrate to affect the rate of cleavage of AdoCbl unless the situation shown in Figure 18 were to prevail. The large isotope effect on formation of the substrate radical slows the overall rate of cleavage of AdoCbl. In a subsequent study [123] Chowdhury and Banerjee measured the temperature dependence of the activation parameters for reaction with [CH3]methylmalonyl-CoA, providing a better estimate of magnitude of the kinetic isotope effect at 49.9. Subsequently computational analysis of the reaction confirmed that the cleavage of AdoCbl was indeed a stepwise process, rather than being concerted with hydrogen atom transfer from substrate to the deoxyadenosyl radical formed on cleavage of AdoCbl [124]. Surprisingly, these computations predicted that the rate constants k1 and k1 in Figure 18 are actually much faster than the rate constant for transfer of the hydrogen atom from the substrate to deoxyadenosine. The equilibrium governing the first step is unfavorable as shown in Figure 18.


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Figure 18. Coupling between cleavage of AdoCbl and formation of the substrate radical in methylmalonyl-CoA mutase. A rapid but unfavorable equilibrium between AdoCbl and the homolytically cleaved dAdo radical and cob(II)alamin precedes a rate-limiting hydrogen atom abstraction from the substrate [124].

Electron paramagnetic resonance has been used to estimate the distance between cob(II)alamin and a succinyl-CoA radical at the active site and their relative orientations [125]. Line broadening induced by heavy atom substitutions in succinyl-CoA indicated that the radical was centered on the carbon a to the free carboxyl. The interspin distance was about 6 A˚ between the two radical centers, and the radical could be modeled in a position very similar to that occupied by succinyl-CoA in a product complex determined by X-ray crystallography. The interspin distance is large for a system that exhibits such large deuterium kinetic isotope effects, which are well above the classical limit and suggest a significant contribution due to hydrogen atom tunneling. In a recent paper, the contribution of hydrogen tunneling to the radical transfer catalyzed by methylmalonyl-CoA mutase has been rigorously analyzed [126]. The authors conclude that the large kinetic isotope effect can only be explained if corner-cutting tunneling decreases the distance over which the system tunnels. Human methylmalonyl-CoA mutase is a mitochondrial enzyme and the only AdoCbl-dependent enzyme in humans, and mitochondrial B12 processing involves reduction of cob(II)alamin to cob(I)alamin, conversion of cob(I)alamin to AdoCbl and then transfer to the methylmalonyl-CoA mutase apoenzyme. Human adenosyltransferase catalyzes the conversion of cob(I)alamin to AdoCbl using ATP as the source of the adenosyl group [127]. However, cob(II)alamin can not be used as the substrate, indicating that the adenosyltransferase does not itself catalyze the reduction of cob(II)alamin to cob(I)alamin. If adenosyltransferase is incubated with cob(II)alamin in the presence of methionine synthase reductase, ATP and NADPH, AdoCbl is formed [128]. Addition of a cob(I)alamin scavenging agent, iodoacetamide, has no effect on this conversion, indicating that the cob(I)alamin is sequestered. The reasonable assumption is therefore that



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reduction occurs while cob(II)alamin is bound to adenosyltransferase. However, we note that methionine synthase reductase is probably not the physiological reducing agent for adenosyltransferase. This reaction occurs in mitochondria, while methionine synthase reductase is cytoplasmic, and patients lacking methionine synthase reductase do not show abnormalities in AdoCbl synthesis. The physiological reducing agent for adenosyltransferase thus remains to be identified. Banerjee and Brunold and their colleagues have shown that adenosyltransferase binds cob(II)alamin in the base-off,His-off form [129], which leads to a more favorable potential for reduction to cob(I)alamin. Furthermore, the cob(II)alamin becomes four-coordinate, lacking water as a ligand, when ATP is present. It is proposed that adenosyltransferase also functions as a chaperone, transferring the base-off AdoCbl to methylmalonyl-CoA mutase [130], which has a higher affinity for the cofactor than adenosyltransferase. While it remains to identify the protein responsible for the reduction of cob(II)alamin to cob(I)alamin in mitochondria, and it remains to determine whether cob(II)alamin is indeed bound to human adenosyltransferase during reduction, the idea of chaperoning this rare and reactive cofactor is highly compelling. The situation should be compared to that in cytoplasmic MetH, where reduction of cob(II)alamin to cob(I)alamin takes place when bound to MetH itself, using electrons from a partner electron transfer protein, and methylation requires AdoMet bound to its own module in methionine synthase. In addition to the reducing agent and adenosyltransferase required for the activity of methylmalonyl-CoA in human mitochondria, a third component, MMAA, is also strongly stimulatory (MMAA is the gene designation for this protein). This protein is a homologue of MeaB, a bacterial protein that is frequently found in operons also containing methylmalonyl-CoA mutase. Mutant strains lacking MeaB are unable to convert methylmalonyl-CoA to succinyl-CoA, although they retain the ability to synthesize AdoCbl [131]. Human patients lacking MMAA belong to the cblA complementation group of patients who present with methylmalonic aciduria [132]. Several studies have been carried out on MeaB from Methylobacterium extorquens, the organism in which MeaB was first studied. MeaB shows homology with GTPases, a family that includes many enzymes involved in assembly of metal cofactors [131]. It forms complexes with holomethylmalonyl-CoA mutase that are enhanced when GTP is bound, and methylmalonyl-CoA mutase stimulates the GTPase activity of MeaB [133]. Furthermore the GTP-bound form of MeaB slows the rate of oxidative inactivation of methylmalonyl-CoA mutase (to form aquacob(III)alamin) by about 15-fold [134]. However, the physiological role of MeaB and its human homologue MMAA in maintaining methylmalonyl-CoA mutase activity remains to be elucidated.


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3.1.4. Isobutyryl-Coenzyme A Mutase Isobutyryl CoA mutase catalyzes the reaction shown in equation (7). This reaction is very similar to the reaction catalyzed by methylmalonyl-CoA

ð7Þ

mutase, with the carboxyl group in methylmalonyl-CoA being replaced by a methyl group in isobutyryl-CoA. Inactivation of the icmA gene in Streptomyces cinnamonensis leads to a strain that is unable to use valine or isobutyryl-CoA as carbon sources [135]. The genes specifying the large and small subunits of isobutyryl-CoA mutase in Streptomyces cinnamonensis have been cloned and sequenced and expressed in E. coli. The icmA gene specifies a 62 kDa large subunit with B40% sequence identity to the large subunits of bacterial methylmalonyl-CoA mutases [136]. However, homologies to the C-terminal cobalamin-binding regions of the latter proteins are lacking. Instead, homologies to the cobalamin-binding regions of methylmalonyl-CoA mutases are found in the icmB gene specifying the 14 kDa small subunit [137]. These homologies include the Asp-X-His-X-X-Gly motif associated with DMB-off,His-on binding of the cofactor. The purified protein is an a2b2 heterodimer. Given the extensive homologies with methylmalonyl-CoA mutase, it is likely that the catalytic mechanisms of these two proteins will be highly similar. Early studies showed that the enzyme catalyzes an intramolecular rearrangement in which the carbonyl thioester of isobutyryl-CoA undergoes a 1,2-migration to the pro-(S) methyl and is replaced by a hydrogen atom at C(3) of n-butyryl-CoA with overall retention [138].

3.1.5. MeaA – A Mutase of Unknown Function A second protein with extensive homology to methylmalonyl-CoA mutase, MeaA, has been identified in Streptomyces collinus and Methylobacterium extorquens. The meaA gene from S. collinus has been cloned and sequenced [139]. It specifies a putative 74 kDa protein with 40% homology to methylmalonyl-CoA mutase and significant homology to the large and small subunits of isobutyryl-CoA mutase. Growth studies suggest that MeaA is Met. Ions Life Sci. 2009, 6, 53–114


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crucial for the production of methylmalonyl-CoA in Streptomyces cinnamonensis, although the actual reaction catalyzed remains to be elucidated.

3.2. Aminomutases and Diol Dehydrases. Isomerization and Elimination Aminomutases catalyze the 1,2 exchange of an amino group with a hydrogen atom, while diol dehydrases catalyze a 1,2 exchange between a hydroxyl group with a hydrogen atom. In a subset of the aminomutase reactions and all the diol dehydrase reactions, the 1,2 exchange is followed by elimination of water. Two enzymes of this type that have been particularly well characterized are diol dehydrase and ethanolamine ammonia lyase, and these will be discussed first. We will then turn to the aminomutases that do not catalyze a subsequent elimination, lysine 2,3-aminomutase and ornithine aminomutase.

3.2.1. Diol Dehydrase Diol dehydrase catalyzes the conversion of (S)-1,2-propanediol (equation 8)

ð8Þ and of (R)-1,2 propanediol (equation 9) to propionaldehyde and water.

ð9Þ While the lack of stereoselection between (S)- and (R)-1,2-propanediols is unusual, the reader will note that the stereochemistry of the two reactions is Met. Ions Life Sci. 2009, 6, 53–114

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MATTHEWS

different [106,140]. As discussed at the beginning of the section on AdoCbldependent mutases, the role of the AdoCbl cofactor as a radical generator was first elucidated for diol dehydrase by observing the exchange of tritium between the 5 0 -position of AdoCbl and the substrate. In contrast to the AdoCbl-dependent enzymes that catalyze carbon skeleton rearrangements, diol dehydrase and ethanolamine ammonia lyase bind AdoCbl in the DMB-on form. This was initially demonstrated by labeling the DMB moiety of AdoCbl with 15N and then determining the EPR spectrum of the enzyme after homolytic cleavage of the carbon-cobalt bond was induced with the suicide substrate 2-methyl-1,2-propanediol [141]. The X-ray structure of diol dehydrase confirmed this conclusion [142]. Diol dehydrase is an a2b2d2 dimer, and the cobalamin is bound at the interface of the a- and b-subunits of each monomer. The a-subunit is an a8b8 barrel, with the cofactor bound at the C-terminal ends of the central b-strands and the substrate 1,2-propanediol bound more deeply in the barrel. The lower face of the cofactor, with its DMB nucleotide coordinated to the cobalt, interacts primarily with the b-subunit. Monovalent cations were reported to be essential cofactors for diol dehydrase [143], and the structure revealed a potassium ion coordinated to the two hydroxyls of the substrate. The distance between the cobalt of the cofactor and C1 and C2 of propanediol were 8.4 A˚ and 9.0 A˚, respectively, in good agreement with data estimating the distance between the radical intermediate observed during steady-state turnover and cob(II)alamin as about 9 A˚ [144]. However, this places the 5 0 -deoxyadenosyl radical that would be generated by homolytic cleavage of AdoCbl too far from the substrate to permit substrate radical formation. Toraya and his colleagues proposed that a simple rotation of the 5 0 -deoxyadenosyl radical around the glycosidic bond would bring C5 0 within 2 A˚ of C1 of the substrate [145], as shown in Figure 19. Subsequent studies clarified the nature of the radical intermediate observed during steady-state turnover of diol dehydrase with 1,2-propanediol. EPR spectroscopy of radicals derived from 13C- and deuterium-labeled substrates established that the radical center resided on C1 [146]. Thus, the intermediate is a substrate-derived radical generated by hydrogen atom abstraction from C1. Further insight into the structure of the radical came from EPR studies of the effects of incorporation of solvent deuterium on the radical signal. These studies indicated that the unpaired electron on the radical center couples with the solvent exchangeable proton on the hydroxyl group at C1 [147]. The latter studies have important implications for the mechanism of hydroxyl migration from C1 to C2, as will be discussed below. The deuterium kinetic isotope effect on kcat observed with [1-2H]-1,2-propanediol is 12, indicating that hydrogen abstraction from C1 of the substrate is rate-limiting in catalysis. However, accumulation of a 5 0 -deoxyadenosyl radical is not seen in either steady-state or stopped-flow analyses. This radical Met. Ions Life Sci. 2009, 6, 53–114


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Figure 19. Mechanism for hydroxyl migration in diol dehydrase. Electron paramagnetic analysis of diol dehydrase reconstituted with 3 0 ,4 0 -anhydroAdoCbl and subjected to homolytic cleavage to form the anhydroadenosyl radical and cob(II)alamin in the presence (A) and absence (B) of (R,S)-1,2-propanediol was used to model the structures. In the absence of substrate, the anhydroribosyl moiety rotates about 601 relative to its position in the presence of substrate, bringing the radical into a position that would be appropiate for hydrogen atom abstraction from substrate and that would not permit formation of a carbon-cobalt bond. Reprinted from [149] with permission from the American Chemical Society, copyright 2006.

would be expected to accumulate if hydrogen abstraction from the substrate were rate limiting. A satisfactory resolution to this dilemma is cartooned in equation (10). The unfavorable equilibrium between AdoCbl and its

ð10Þ

homolytic cleavage products prevents the accumulation of observable 5 0 deoxyadenosyl radical, and leads to the observed kinetic isotope effect on the accumulation of the substrate-derived radical intermediate in the rate-limiting step of catalysis. Also consistent with this proposal is the observation that detectable formation of cob(II)alamin requires the presence of substrate [148]. Support for this postulate also comes from studies with diol dehydrase reconstituted with 3 0 ,4 0 -anhydroAdoCbl (Figure 20). The introduction of a double bond adjacent to the radical center generated by homolytic cleavage of Met. Ions Life Sci. 2009, 6, 53–114

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Figure 20.

MATTHEWS

Structure of 3 0 ,4 0 -anhydroadenosylcobalamin.

this cofactor analogue greatly stabilizes the resulting radical, and results in its accumulation during reaction with 1,2-propanediol [149]. In fact, homolytic cleavage of 3 0 -4 0 -anhydroAdoCbl bound to diol dehydrase is also observed in the absence of substrate. Still to be resolved is the mechanism of hydroxyl migration from C1 to C2 following hydrogen atom abstraction from C1 of 1,2-propanediol. A variety of mechanisms have been proposed (Figure 21) involving (a) general base catalysis, (b) general acid catalysis, (c) partial protonation (hydrogen bonding), (d) electrophilic catalysis by the activating potassium ion, or (e) combined general acid/base (push-pull) catalysis (discussed in [147]). The demonstration that the proton remains on the C1 hydroxyl group of the substrate-derived radical argues against the base-catalyzed mechanism for rearrangement [147]. Activation of the enzyme by thallous ion instead of K1, which introduces a spin 1/2 metal in close proximity to the substrate radical, fails to lead to spin coupling with the substrate-derived radical, also disfavoring a role for K1 in catalyzing the hydroxyl migration [147]. His143 is in hydrogen bonding distance to the hydroxyl on C2 of the substrate and Met. Ions Life Sci. 2009, 6, 53–114


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Figure 21. Proposed mechanisms for hydroxyl migration in diol dehydrase. The mechanisms proposed involve (a) general base catalysis, (b) general acid catalysis, (c) partial protonation (hydrogen bonding), (d) electrophilic catalysis by the activating potassium ion, or (e) combined general acid/base (push-pull) catalysis. Reprinted from Schwartz et al. [147] with permission from Protein Science, copyright 2007. Met. Ions Life Sci. 2009, 6, 53–114

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Glu170 is in hydrogen bonding distance to the hydroxyl on C1. Mutation of these residues to alanine reduces kcat by 77-fold and 38,000-fold respectively [150], making the push-pull mechanism shown in (e) of Figure 21 highly attractive. What then is the role of the essential monocation activator? Recent studies have shown that K1 activates the spontaneous cleavage of AdoCbl in the absence of substrate [151]. The reaction is observed as the formation of aquacob(III)alamin and requires both O2 and K1. These studies suggest that the binding energy of K1 is used to enforce a conformational change in the enzyme that strains the carbon-cobalt bond of AdoCbl, accelerating cleavage. This conclusion is in agreement with conclusions drawn from crystallographic studies of diol dehydrase with the coenzyme analogue adeninylpentylcobalamin bound [152]. This analogue has an increased distance between the adenosine and the cobalamin. These studies suggested that in diol dehydrase with K1 bound, the AdoCbl would be bound in a strained conformation even in the absence of substrate. Spontaneous cleavage of AdoCbl bound to diol dehydrase would result in inactivation in the cellular milieu, which contains both oxygen and potassium ions, because the aquacobalamin is extremely tightly bound and this form of the enzyme is inactive. Mori and Toraya [153] have identified a reactivating factor, a chaperone-like protein with ATPase activity. In the presence of ADP, the chaperone mediates the release of aquacobalamin from the inactive enzyme and binds tightly to the apoenzyme, while in the presence of ATP, the apoenzyme is released and can be reconstituted with AdoCbl. This chaperone comprises two subunits, gene products of the ddrA and ddrB genes in Klebsiella oxytoca.

3.2.2. Ethanolamine Ammonia Lyase Ethanolamine ammonia lyase catalyzes the conversion of ethanolamine to acetaldehyde and ammonia. The reaction proceeds as shown in equation (11). The enzyme is an a6b6 oligomer, and recent studies suggest that a mol

ð11Þ

of AdoCbl is bound per ab protomer [154]. The predicted masses of the aand b-subunits from E. coli are 50 and 32 kDa, respectively. Although the subunits show little homology with those of other B12-dependent enzymes, Met. Ions Life Sci. 2009, 6, 53–114


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the larger subunit does show limited homology with the subunits of methylmalonyl-CoA mutase from P. shermanii [154], and recent modeling of the large subunit of the enzyme from Salmonella typhimurium suggests that this subunit is indeed a b8a8 barrel [155]. By analogy with diol dehydrase, we might expect the AdoCbl to be bound at the interface between the small and large subunits. As in diol dehydrase, the AdoCbl is bound in the ‘‘DMB-on’’ form [156]. The X-ray structure of ethanolamine ammonia lyase has not been determined, but sophisticated spectroscopic studies have revealed its mechanistic similarity with diol dehydrase and also provided some unique insights into the catalytic mechanism. One of the first issues was whether hydrogen transfer actually occurred directly between the AdoCbl radical and the substrate, or whether an intermediary such as a protein radical might be involved. Following cleavage of AdoCbl in the presence of substrate, the substrate-derived radical was found to be more than 10 A˚ from the unpaired electron of cob(II)alamin [157]. In a landmark study, electron nuclear double resonance and specific 2H- and 13C-labeling of the substrate was used to show that the methyl group of 5 0 -dAdo was positioned 3.4 A˚ from C1 0 of the substrate radical, in a perfect position to mediate direct hydrogen transfer between cofactor and substrate [158]. Similar conclusions were reached using pulsed EPR [159]. Thus, the 5 0 -carbon of AdoCbl migrates B7 A˚ from its position when the cofactor is intact, to its position when the substrate radical is formed. One major difference between diol dehydrase and ethanolamine ammonia lyase is that, while the stereochemistry of the diol dehydrase reaction requires that the hydroxyl group migrates from C1 to C2 prior to elimination of water, there is no such compelling evidence that the amino group of ethanolamine migrates rather than simply undergoing elimination from C2 after hydrogen atom abstraction at C1. However, arguing by analogy with all the other AdoCbl-dependent isomerases, we might expect that such migration does actually occur. A second difference is the apparent absence of a requirement for a cation (potassium ion or ammonia) for catalysis. In the modeling of the active site of the Salmonella enzyme, Sun and Warncke suggest that the guanidinium side chain of Arg160 occupies the same position as the potassium ion in the active site of diol dehydrase [155].

3.2.3. Lysine 5,6-Aminomutase Lysine 5,6-aminomutase was initially characterized in the laboratory of Theresa Stadtman, and was shown to be an AdoCbl-dependent enzyme that catalyzed a 1,2-migration of the e-amino group of D-lysine with concomitant Met. Ions Life Sci. 2009, 6, 53–114

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reverse migration of a hydrogen atom, as shown in equation (12) [160]. The stereochemistry of the reaction has not yet been determined, so we do

ð12Þ

not yet know which of the two hydrogens on the d-carbon of D-lysine migrates. More recently, the enzyme was cloned, sequenced, expressed and purified from Clostridium sticklandii [161]. In agreement with earlier studies, in which the enzyme was isolated as a 170 kDa complex of 55 and 30 kDa subunits, the enzyme is an a2b2 heterodimer composed of 57 kDa a-subunits and 29 kDa b-subunits. The small subunit contains the Asp-X-His-X-X-Gly sequence characteristic of His-on binding of the AdoCbl cofactor. The substrate D-lysine forms an external aldimine linkage (Figure 22) with the pyridoxal phosphate cofactor that is required for enzyme activity [162]. The pyridoxal phosphate is proposed to stabilize radical intermediates formed following hydrogen atom abstraction from the substrate as shown in Figure 22 [163]. The X-ray structure of the enzyme [164] revealed that the large subunit is an a8b8 barrel and positions the pyridoxal phosphate cofactor at the Cterminal end of the barrel strands. The crystal structure was obtained in the absence of D-lysine, and revealed that the pyridoxal phosphate was linked as

Figure 22. Proposed role for pyridoxal phosphate in the reaction catalyzed by lysine 5,6-aminomutase [163]. Met. Ions Life Sci. 2009, 6, 53–114


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an internal aldimine with Lys144 of the small subunit. The AdoCbl is sandwiched between the large and small subunits, in the predicted DMB-off,His-on conformation. However, the AdoCbl is far from the pyridoxal phosphate binding site in this ‘‘resting’’ conformation of the enzyme. The covalent bond between pyridoxal phosphate and Lys144b locks the two subunits together in a conformation that leaves the active site accessible to solvent and substrate and prevents cleavage of AdoCbl. The authors propose that substrate binding will lead to a rearrangement that will sequester the active site and position AdoCbl appropriately for catalysis.

3.2.4. Ornithine 4,5-Aminomutase This enzyme catalyzes the reaction shown in equation (13). Until very recently, the enzymatic reaction had only been studied in crude extracts,

ð13Þ

where the product was shown to be (2R,4S)-diaminopentanoate. However the stereochemistry of the migrating hydrogen atom in ornithine has not been determined. In 2001, the oraE and oraS genes specifying the large and small subunits of D-ornithine amino mutase were cloned and sequenced from Clostridium sticklandii [165], and in 2004, the enzyme was successfully expressed in E. coli and purified to homogeneity [166]. Despite the lack of sequence homology with other AdoCbl-dependent enzymes, the properties of the purified ornithine aminomutase are very similar to those of lysine 5,6aminomutase. The enzyme is an a2b2 heterodimer, and requires pyridoxal phosphate for activity in addition to AdoCbl.

3.3. Adenosylcobalamin-Dependent Ribonucleotide Triphosphate Reductase Ribonucleotide triphosphate reductase catalyzes the reaction shown in equation (14), which results in oxidation of two active site thiols to form a Met. Ions Life Sci. 2009, 6, 53–114

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MATTHEWS

ð14Þ disulfide. Regeneration of the enzyme by reduction of the disulfide bond is required for sustained turnover, and is accomplished by a series of electron transfers from NADPH to thioredoxin reductase, to thioredoxin, and then to ribonucleotide triphosphate reductase as shown in equation (15). S E S

thioredoxin + NADPH thioredoxin reductase

SH E

+ NADP+

ð15Þ

SH

Enzyme catalysis requires AdoCbl, which serves as a radical generator as it does in other AdoCbl-dependent enzymes. However, a unique feature of ribonucleotide triphosphate reductase is that hydrogen atom abstraction from the substrate is not catalyzed by the 5 0 -deoxyadenosyl radical, but rather by a thiyl radical generated by hydrogen atom abstraction from a cysteine residue on the protein. The enzyme from Lactobacillus leichmanii has been most extensively studied. In contrast to the intramolecular tritium transfer seen in other AdoCbldependent enzymes, if ribonucleotide triphosphate reductase labeled with tritium in the 5 0 position of AdoCbl is incubated with the allosteric activator dGTP and reductant, the label is quantitatively transferred to solvent [167]. Stubbe and her colleagues then showed using single turnover experiments that ribonucleotide triphosphate reductase catalyzes the cleavage of the 3 0 -carbon-hydrogen bond of UTP with quantitative release of label to solvent when the 3 0 -position is tritiated [168]. The authors proposed a working hypothesis for the enzyme that involved the generation of a protein radical that in turn generated the substrate radical by direct hydrogen atom abstraction of the 3 0 -hydrogen of UTP. Cysteine 408 was subsequently shown to be the site of formation of the protein radical [169], and rapid freeze quench electron paramagnetic resonance of an intermediate in the exchange of the 5 0 -hydrogens of AdoCbl with solvent catalyzed by ribonucleotide triphosphate reductase containing specifically deuterated cysteines demonstrated a cysteine-centered radical that was spin-coupled to cob(II)alamin. These studies also demonstrated that the formation and decay of the cysteine-based radical signal proceeded at rates consistent with its involvement in catalysis. The detailed mechanism proposed for the catalytic cycle [170] is shown in Figure 23. Met. Ions Life Sci. 2009, 6, 53–114


Figure 23.

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Proposed mechanism for AdoCbl-dependent ribonucleotide reductase [170].

Following the cloning and sequencing of ribonucleotide triphosphate reductase from L. leichmanii [171], a crystal structure of the enzyme was obtained with the AdoCbl analogue adeninylpentylcobalamin bound [172]. The monomeric enzyme folds as a ten stranded a/b barrel with Cys408 Met. Ions Life Sci. 2009, 6, 53–114

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positioned at the tip of a hairpin loop at the bottom of the barrel. The AdoCbl analogue is bound with DMB in the ‘‘base-on’’ conformation, as previously predicted [173]. One issue that has recently been clarified concerns the mechanism of formation of the thiyl radical, which could be generated in a stepwise fashion by cleavage of AdoCbl to form 5 0 -deoxyadenosyl radical and cob(II)alamin, or in a concerted fashion in which a 5 0 -deoxyadenosyl radical is not an intermediate. When ribonucleotide reductase was incubated with stereoselectively deuterated (5 0 R)-[5 0 -2H]AdoCbl and the allosteric activator dGTP, the 5 0 deuterium was stereochemically scrambled [174]. This scrambling occurred in both the wild-type enzyme and the Cys408Ala mutant, even though exchange of deuterium with solvent did not occur in the mutant enzyme. The implication is that transient cleavage of the Co-C5 0 bond of AdoCbl occurs in the presence of the allosteric activator even in the absence of Cys408. The second mechanistic issue concerns the sequence by which the activesite disulfide is reduced at the end of each turnover. As previously mentioned, the electrons ultimately come from NADPH, and are transferred to the redox active disulfide of thioredoxin by thioredoxin reductase. Reduction of ribonucleotide triphosphate reductase by thioredoxin requires two cysteines at the C-terminus of the protein, Cys731 and 736 [169], and these two cysteines are thought to shuttle reducing equivalents between reduced thioredoxin and the active site disulfide. This C-terminal extension is disordered in the crystal structure of the enzyme [172].

4. CONCLUDING REMARKS The author hopes that this review highlights the explosion of information recently obtained about the growing family of characterized corrinoiddependent enzymes, and particularly the mounting insights available from structural analyses of these proteins. We now appreciate the vital role that corrinoid-dependent methyltransferases play in central metabolic pathways in prokaryotes and Archaea. A challenge for the future will be to reach a detailed understanding of how the membrane-associated energy conserving corrinoid methyl transferase couples sodium ion transport with methyl transfer from methylcobalamin to the thiol of coenzyme M. The elegant spectroscopies that have been applied to the AdoCbldependent enzymes have greatly clarified the nature of the radical intermediates and structural analyses have revealed the type of molecular motions required to allow hydrogen atom transfer from substrate to the deoxyadenosyl radical and back to product. These complicated enzymes have posed a challenge to enzymologists and chemists for the past 70 years, Met. Ions Life Sci. 2009, 6, 53–114


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and it has been thrilling to review the elegant experiments that step-by-step revealed the details of catalysis.

ACKNOWLEDGMENTS Research in the author’s laboratory is funded by National Institutes of Health Grant GM24908.

ABBREVIATIONS AND DEFINITIONS AdoCbl AdoMet ADP ATP CoA dGTP DMB EPR EXAFS GTP Hcy MAP MetH NADP(H) NMR UroD UTP

adenosylcobalamin S-adenosylmethionine adenosine 5 0 -diphosphate adenosine 5 0 -triphosphate coenzyme A deoxyguanosine 5 0 -triphosphate dimethylbenzimidazole electron paramagnetic resonance extended X-ray absorption fine structure guanosine 5 0 -triphosphate homocysteine methyltransferase-activating protein cobalamin-dependent methionine synthase from E. coli nicotinamide adenine dinucleotide phosphate (reduced) nuclear magnetic resonance uroporphyrinogen decarboxylase uridine 5 0 -triphosphate

REFERENCES 1. S. W. Ragsdale, P. A. Lindahl and E. Munck, J. Biol. Chem., 1987, 262, 14289–14297. 2. E. Stupperich, H. J. Eisenger and S. P. J. Albracht, Eur. J. Biochem., 1990, 193, 105–109. 3. S. W. Ragsdale, The Acetogenic Corrinoid Proteins, in Chemistry and Biochemistry of B12, Ed. R. Banerjee, John Wiley & Sons, New York, 1999. 4. R. V. Banerjee, N. L. Johnston, J. K. Sobeski, P. Datta and R. G. Matthews, J. Biol. Chem., 1989, 264, 13888–13895.


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Met. Ions Life Sci. 2009, 6, 115–132

3 Nickel-Alkyl Bond Formation in the Active Site of Methyl-Coenzyme M Reductase Bernhard Jaun a and Rudolf K. Thauer b a

Organic Chemistry ETHZ, ETH Ho¨nggerberg HCI E317, CH-8093 Zu¨rich, Switzerland <[email protected]> b Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Strasse, D-35043 Marburg, Germany

ABSTRACT 116 1. INTRODUCTION 116 2. NICKEL-CARBON BOND FORMATION IN FREE 119 COENZYME F430 3. NICKEL-ALKYL BOND FORMATION IN MCR UPON INACTIVATION WITH ALKYL HALIDES 120 3.1. 3-Sulfonatopropyl-Ni(III)F430 Formation by Reaction of 120 MCRred1 with 3-Bromopropane Sulfonate 3.2. Methyl-Ni(III)F430 Formation in MCRred1 by Reaction with Methyl Bromide 122 4. METHYL-NICKEL BOND FORMATION IN METHYLCOENZYME M REDUCTASE DURING CATALYSIS? 123 4.1. Methylation of Ni(I)F430 in the Active Site via Nucleophilic Substitution 125 4.2. Methylation of Ni(I)F430 in the Active Site via Oxidative Addition 127 4.3. Catalytic Mechanism of Methyl-Coenzyme M Reductase Not Involving Metal-Carbon Bond Formation 128


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5. OBSERVATIONS TO BE FOLLOWED UP ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

129 129 129 130

ABSTRACT: Methyl-coenzyme M reductase (MCR) catalyzes the methane-forming step in methanogenic archaea and most probably also the methane-oxidizing step in methanotrophic archaea. The enzyme contains coenzyme F430 as prosthetic group. F430 is a nickel porphinoid that has to be in the reduced Ni(I) state for the enzyme to be active. The presently discussed catalytic mechanisms of MCR can in principle be divided into two basic models. In one model the key intermediate features a methyl-Ni(III) species being either formed in a nucleophilic substitution reaction or in an oxidative addition reaction. In the other model first the thioether sulfur of methyl-coenzyme M binds to the Ni(I), which subsequently results in the release of the methyl group as methyl radical leaving behind a Ni(II)-sulfur bond. The experimental evidence for and against a methyl-nickel intermediate is reviewed. KEYWORDS: anaerobic oxidation of methane catalytic mechanism of methane formation coenzyme F430 methane formation methyl-coenzyme M reductase methylnickel intermediate nickel enzymes nickel porphinoid

1. INTRODUCTION Methyl-coenzyme M reductase (MCR) is a nickel enzyme found in relatively high concentrations in all methanogenic and methanotrophic archaea [1–4]. The enzyme catalyzes the reversible reaction of methyl-coenzyme M (CH3-S-CoM; 2-(methylthio)ethanesulfonate) with coenzyme B (CoBSH; N-7-thioheptanoyl-O-phospho-L-threonine) to methane and the heterodisulfide (CoM-S-S-CoB) of coenzyme M (CoM-SH) and coenzyme B (reaction 1) at a specific rate of approximately 100 mmol methane formed per min and per mg protein which is equivalent to a turnover number of 250 s1 [5]. CH3 -S-CoMþCoB-SHÐ CH4 þ CoM-S-S-CoB 0

DG0 ¼ 30 kJ=mol

ð1Þ

MCR has a molecular mass of 300 kDa and is composed of three different subunits in an a2b2g2 arrangement, which form two structurally and probably also functionally coupled active sites, each containing one coenzyme F430 as prosthetic group. Coenzyme F430 is a nickel hydroporphyrin of Met. Ions Life Sci. 2009, 6, 115–132

METHYL-COENZYME M REDUCTASE: Ni-C BOND FORMATION

117

Figure 1. Structure of coenzyme F430 with its nickel axially coordinated by the carboxamide group of glutamine 147 in the active site of methyl-coenzyme M reductase from Methanothermobacter marburgensis. View onto the b-face of the corphin ring. Also shown are the structures of the substrates methyl-coenzyme M and of coenzyme B.

unique structure (Figure 1), corphin being the name proposed by Eschenmoser for this class of tetrapyrroles [6]. F430 has to be in the reduced Ni(I) state for the enzyme to be active and it has been proposed that also the Ni(III) state is involved in the catalytic cycle [7–9]. The pentamethyl ester of coenzyme F430 (F430M) shows a redox potential Eo 0 of nearly 0.6 V for the Ni(II)F430M/Ni(I)F430M couple (in acetonitrile) [10] and one of nearly +1.6 V for the Ni(III)F430M/Ni(II)F430M couple (in acetonitrile) [11]. Compared to the normal hydrogen electrode (NHE) the first redox potential is lower than that of the hydrogen electrode in water at pH 7 (0.42 V) and the second redox potential is higher than that of the oxygen electrode in water at pH 7 (0.81 V) explaining why only Ni(II)F430 is stable in water. Ni(II)F430 can be reduced to Ni(I)F430 in water at a mid potential of approximately 600 mV but only if the proton concentration is kept below pH 7. Consistently, in MCR the nickel corphin is buried deeply Met. Ions Life Sci. 2009, 6, 115–132

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within the protein in a water free hydrophobic pocket accessible from the outside only via a narrow 50 A˚ long channel as revealed by the crystal structures of inactive Ni(II) forms [12–15]. The redox potentials of F430 in its MCR-bound form are not known. However, that of the Ni(II)F430/Ni(I)F430 couple is probably very close to that of the free cofactor as deduced from the similar UV-visible spectra of free and MCR-bound Ni(II)F430 and of the similar UV-visible and EPR spectra of free- and MCR-bound Ni(I)F430. In contrast, the redox potential of the Ni(III)F430/Ni(II)F430 couple of free- and MCR-bound F430 are predicted to be significantly different. Neither the UV-visible nor the EPR spectrum of free Ni(III)F430M show similarity with the spectra of the different MCR-bound forms of coenzyme F430 that are formally in the Ni(III) valence state [1]. Active MCR is referred to as MCRred1 and its Ni(I)-derived EPR signal as the MCRred1 signal. In Ni(I)F430 and in MCRred1 the unpaired electron resides mainly in the dx2y2 orbital, the nickel dz2 orbital being filled with two electrons. MCRred1 can be converted to other EPR active but enzymatically inactive forms which are distinguished via differences in their Ni(I)-[d9] or Ni(III)-[d7] EPR spectra: MCRred1/red2 (Ni(I)) by reaction of MCRred1 with coenzyme M and coenzyme B (reaction 2) [16,17]; MCRox1(Ni(III)) by reaction of MCRred1/2 with polysulfide (reaction 3) [18]; MCRox1 back to MCRred1 by reduction with titanium(III)citrate [Ti(III)] (reaction 4) [5]; MCRPS (Ni(III)) by reaction of MCRred1 with 3-bromopropane sulfonate (BPS) (reaction 5) [19]; MCRPS back to MCRred1 by reaction with sulfide (reaction 6) [20]; MCRMe(Ni(III)) by reaction of MCRred1 with methyl bromide (BrMe) or methyl iodide (reaction 7) [21,22]; and MCRMe back to MCRred1 by reaction with coenzyme M (reaction 8) [22]. (In the literature MCRPS is also referred to as MCRBPS and MCRMe as MCRBrMe). MCRred1 þ CoM-SH þ CoB-SH Ð MCRred1=2 MCRred1=2 þ polysulfide ! MCRox1 MCRox1 þ TiðIIIÞ ! MCRred1

ðat pH 7Þ

ðat pH 7Þ

ð3Þ

ðat pH 10Þ

MCRred1 þ 3-bromopropane sulfonate ! MCRPS þ Br

ð2Þ

ð4Þ ðat pH 7Þ

ð5Þ

MCRPS þ S2 !MCRred1 þ 3-mercaptopropane sulfonate ðat pH 10Þ

ð6Þ

MCRred1 þ BrCH3 ! MCRMe þ Br

ð7Þ

Met. Ions Life Sci. 2009, 6, 115–132

ðat pH 7Þ


119

MCRMe þ CoM-SH ! MCRred1 þ CH3 -S-CoM

ð8Þ

ðat pH 7Þ

In the presence of coenzyme M (which is an inhibitor of MCR) and coenzyme B maximally 50% of the MCRred1 signal is converted into the MCRred2 signal indicating half-of-the-sites reactivity [17]. The MCRred2 signal is composed of an axial component (MCRred2a) and a strongly orthorhombic component (MCRred2r). The two forms are in a temperature dependent equilibrium with MCRred1 [23]. In MCRred1, MCRox1, MCRPS and MCRMe the nickel of F430 is axially ligated from the a-face (Figure 1) by the oxygen of the carboxamide group of glutamine147a,a0 [1,9]. This ligand is displaced in MCRred2 [24]. The upper axial ligand in MCRox1 is the sulfur of the thiolate group of coenzyme M [25], in MCRPS it is a 3-sulfonatopropyl group [26] and in MCRMe a methyl group [21,22]. In MCRred2a it is a hydride and in MCRred2r a hydrogen and a sulfur, presumably as the result on an oxidative addition of the S-H bond of coenzyme M to Ni(I)F430 giving a (S)(H)Ni(III)F430 species [24]. In MCRred1 the b-axial coordination site appears to be unoccupied [27]. The properties on MCR and of its nickel corphin cofactor coenzyme F430 have been extensively reviewed by the authors two years ago in Volume 2 of ‘‘Metal Ions in Life Sciences’’ [1]. This allows the present chapter to completely focus on what is known about metal-carbon bonds in MCR and its cofactor.

2. NICKEL-CARBON BOND FORMATION IN FREE COENZYME F430 Ni(II)F430M (M for pentamethyl ester) in dry dichloromethane, chloroform or acetonitrile is in a diamagnetic d8 low spin state. In this state the nickel in F430 is square planar tetracoordinated, but has a pronounced tendency to bind additional ligands in the axial positions such as chloride (log K1 ¼ 5.4; log K2 ¼ 3.7), imidazole (log K1 ¼ 2.7; log K2 ¼ 2.2) or water. Penta- and hexacoordination is associated with a change from low spin to high spin, a shift of the absorbance band at 422 nm (e ¼ 21 mM1cm1) by a few nm and a gain in band intensity [1]. In water Ni(II)F430 has an absorbance maximum at 430 nm and an extinction coefficient of 23 mM1cm1. When methyl iodide is allowed to react with Ni(I)F430M in a 1:2 stoichiometric ratio at low temperature, the color of the solution changes from green Ni(I)F430M to brown-orange within seconds without methane being formed. When acid is added, the color changes to that of Ni(II)F430M and more than 80% of the theoretical amount of methane is generated. Addition of deuterated acid gives over 85% CH3D. This experiment proves that an Met. Ions Life Sci. 2009, 6, 115–132

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intermediate is formed, which can be dissociated to Ni(II)F430M and methane by protonation [28–30]. Its properties are consistent with those expected for a methyl-Ni(II) derivative with an axial nickel-carbon bond. CD3-Ni(II)F430M and CH3-Ni(II)F430M were independently generated at low temperature by stoichiometric reaction with (CD3)2Mg and (CH3)2Mg, respectively, and their structure was verified by 2H and 1H NMR spectroscopy [30]. The dissociation energy for the Ni(II)-C bond was estimated to be of the order of 80 kJ/mol [31]. Formation of methyl-Ni(III)F430M as an intermediate is discussed for the reaction of Ni(I)F430M with methyl iodide to give methyl-Ni(II)F430M. However, all attempts to demonstrate the existence of this compound outside the enzyme have failed until now although theoretical modeling (DFT) predicts that methyl-Ni(III) should be stable [32]. One reason probably is that any methyl-Ni(III)F430M formed will immediately react with excess Ni(I)F430M to methyl-Ni(II)F430M and Ni(II)F430M. The redox potential of the methyl-Ni(III)F430M/methyl-Ni(II)F430M couple has been estimated to be near +0.45 V [1] which is one Volt more positive than the redox potential E1 0 ¼0.6 V of the Ni(II)F430M/Ni(I)F430M couple [10]. Coenzyme F430 has been shown to catalyze the reductive dehalogenation of CCl4, CHCl3, CH2Cl2, and CH3Cl with Ti(III) in aqueous solution. Trichloromethyl-Ni(III)F430, dichloromethyl-Ni(III)F430, monochloromethyl-Ni(III)F430, and methyl-Ni(III)F430, respectively, have been proposed to be likely intermediates [33]. The reaction of Ni(I)-octaethylisobacteriochlorin (a structural cousin of F430) with alkyl halides has been investigated in detail and has been interpreted as proceeding via alkyl-Ni(III) species that undergo reduction to the alkyl-Ni(II), followed by protonolysis yielding the alkane [34].

3. NICKEL-ALKYL BOND FORMATION IN MCR UPON INACTIVATION WITH ALKYL HALIDES 3.1. 3-Sulfonatopropyl-Ni(III)F430 Formation by Reaction of MCRred1 with 3-Bromopropane Sulfonate 3-Bromopropane sulfonate is the most potent inhibitor of MCR known to date. The methyl-coenzyme M analogue binds in the active site of MCRred1 with a nanomolar inhibition constant (Ki), where it reacts with the Ni(I) of F430 to give MCRPS which has a nickel based, axial X-band continuouswave (CW) EPR spectrum [19]. In view of the known reactivity of free Ni(I)F430 towards alkyl halides [33] and the results of DFT calculations [32] the possibility was considered that the MCRPS species might Met. Ions Life Sci. 2009, 6, 115–132


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correspond to an alkyl-Ni(III)F430 derivative resulting from reaction of Ni(I)F430 with BPS to give a bromide ion and O3S(CH2)3-Ni(III)F430 in the active site of MCR through what is formally an oxidative addition of BPS to Ni(I):

O3 SðCH2 Þ3 Br þ NiðIÞF430 ! O3 SðCH2 Þ3 -NiðIIIÞF430 þ Br

ð9Þ

To test this hypothesis, Hinderberger et al. [26] synthesized [3-13C]-3bromopropane sulfonate, then treated the active enzyme with it, and investigated the resulting 13C-MCRPS samples by using high frequency CWEPR (W-band, microwave frequency ca. 94 GHz), high-resolution pulse electron nuclear double resonance (ENDOR), and hyperfine sublevel correlation (HYSCORE) spectroscopy. As a control, MCRPS samples induced by reaction of MCRred1 with unlabeled 3-bromopropane sulfonate were also studied. To further clarify the coordination geometry, the proton signals derived from the two g protons and the two b protons of the propane sulfonate moiety were investigated. The interpretation of the spectroscopic results in terms of the binding situation in MCRPS is shown in Figure 2A. In MCRPS there is a bond between the nickel center of F430 and the Cg atom of the propane sulfonate residue. Approximately 7% of the spin

Figure 2. Proposed coordination of the nickel in the active site of MCR after reaction of MCRred1 with 3-bromopropane sulfonate to form MCRPS. (A) The pz orbital of the proposed near sp2-hybrid carbon atom has been drawn to indicate the direction of the Ni-Cg bond. The asterix marks the position of 13C labeling. Regarding the detailed ligation of Ni see Figure 1. (B) Newman projection along the Cg-Cb bonding. For clarity the Ni-Cg has been elongated, the pz orbital of Cg has been drawn in gray and the g protons have been omitted. R ¼ -CH2-SO 3 . Reproduced from Hinderberger et al. [26] with permission of Wiley-VCH, copyright (2006). Met. Ions Life Sci. 2009, 6, 115–132

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population resides on the Cg atom and 75% in the Ni dx2y2 orbital in close agreement with the results of DFT calculations on a CH3-Ni(III)F430 model structure [32]. The Ni-Cg bond is slightly tilted (ca. 201) away from the gz principle axis direction. Hybridization of the Cg atom is closer to sp2 than sp3 resulting in a flattened bonding geometry at Cg: the two protons Hg1 and Hg2 are situated in (or close to) a nodal plane of the nonhybridized pz orbital of the Cg atom that hence has very little overlap with the hydrogen 1s orbitals [26]. The Newman projection of the conformation at the Cb atom as deduced from proton coupling is shown in Figure 2B. It was noted by the authors that the EPR data do not allow the determination of the total number of electrons in the system, and that therefore the assignment of a formal alkyl-Ni(III) state rests on the assumption that after the reaction of MCRred1 with 3-bromopropane sulfonate a bromide ion is generated as a reaction product [26]. In the meantime it was found that upon acid denaturation of MCRPS the C-Ni bond is protonolized yielding propane sulfonate [35]. This product was also identified when MCRPS was re-reduced with Ti(III) to MCRred1. MCRPS can also be converted to active MCRred1 by treatment with sodium borohydride [20]. Upon addition of thiols to MCRPS at pH 10 the active MCRred1 state is regenerated with the concomitant formation of the respective thioether [20,35,36]. All these findings point to the presence of an alkyl-Ni(III) bond in MCRPS. MCRred1 was found to react with 4-bromobutyrate and other o-brominated carboxylic acids with carbon chain length up to 16. All of these compounds give rise to an alkyl-Ni intermediate with an EPR signal similar to that of the MCRPS species. Reaction of the alkyl-Ni adduct, formed from brominated acids with eight or fewer total carbons, with CoM-SH as nucleophile at pH 10 results in the formation of a thioether coupled to the regeneration of the active MCRred1 state [22,36]. Reactivation is highest with the smallest free thiol HS–. Interestingly, MCRPS can also be reductively activated with analogues of HS-CoB such as HS-CoB8 (N-8-thiooctanoyl-O-phospho-L-threonine) and HS-CoB9 (N-9-thiononanoyl-O-phospho-L-threonine) but not with coenzyme B (N-7-thioheptanoyl-O-phospho-L-threonine) [20]. MCRred1 alkylated with 4-bromobutyrate was surprisingly found to undergo ‘‘self-reactivation’’ at pH 10 to generate the MCRred1 state and the ester between 4-bromobutyrate and 4-hydroxybutyrate [36].

3.2. Methyl-Ni(III)F430 Formation in MCRred1 by Reaction with Methyl Bromide The active enzyme MCRred1 reacts rapidly and irreversibly with simple alkyl halides such as chloroform. The latter has been routinely used to quench the Met. Ions Life Sci. 2009, 6, 115–132


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Ni(I) state yielding an EPR silent MCR. Also the inhibitor 2-bromoethanesulfonate (BES) quenches the Ni(I) state with the concomitant formation of a protein based radical [19]. To the contrary, reaction of BPS with MCRred1 yields MCRPS with an alkyl-Ni(III) bond. Apparently it needs a methyl-coenzyme M related structure for the alkyl-Ni(III) derivative to be stable. It was therefore surprising that upon reaction of methyl bromide with MCRred1 the enzyme was converted into a form MCRMe with an UV-visible and EPR spectrum very similar to that of MCRPS [21,22]. Complete conversion of the MCRred1 signal into the MCRMe signal was achieved by incubation of MCRred1 in 50 mM Tris/HCl pH 9.0 with an approximately 50 fold excess of methyl bromide or methyl iodide (added as saturated aqueous solution). The signal decays to an EPR silent form with a t1/2 of 20 min at room temperature. Continuous wave and pulse electron nuclear double resonance and hyperfine sublevel correlation spectroscopy of the enzyme alkylated with CH3Br, 13CH3Br or CD3Br (or the respective iodides) revealed that the methyl group from the methyl halide becomes directly bound to the nickel ion after reaction. From the EPR data it was calculated that the nickel has a spin population of 81% which resides in the Ni(III) dx2y2 orbital. The most reasonable picture of the H3C-Ni(III) coordination is via an interaction of the filled nickel dz2 orbital with the empty orbital from the cation CH+ 3 , indicating that the halogen atom is lost as halogenide ion [21,22]. When the CH3-Ni(III) species was reacted with CoM-SH, active MCRred1 was regenerated with a rate constant of 0.044 s1 forming methyl-coenzyme M as demonstrated by mass spectrometry [22]. The puzzling observation that the Ni(I) ion of MCRred1 and the Ni(III) ion of MCRMe and MCRPS both have dx2y2 ground configurations, even though the formal electron count on Ni differs by two, is made understandable through the cartoon bonding scheme for methyl-Ni(III) in Figure 3 [22]. The scheme explains why the odd-electron orbital is largely unperturbed by the oxidative addition reaction and why the transfer of spin to CH3 is minimal: the Ni-C two electron bond does not involve the odd electron, which in effect is a spectator to the reaction! The change in g-values can be assigned primarily to changes in the d-orbital splitting [22].

4. METHYL-NICKEL BOND FORMATION IN METHYLCOENZYME M REDUCTASE DURING CATALYSIS? Different catalytic mechanisms are presently discussed for the reaction of methyl-coenzyme M and coenzyme B to methane and the heterodisulfide (reaction 1). They are based on density function theory calculations, on the Met. Ions Life Sci. 2009, 6, 115–132

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Figure 3. Cartoon bonding scheme for the Ni–C fragment obtained after oxidative addition of the Br-C bond of methyl bromide to Ni(I) in MCRred1. The scheme shows the highest two d-orbitals of the parent Ni(I) center, occupied by three electrons, and the empty spn hybrid of the hypothetical CH+ 3 , with which this center reacts to form the CH3-Ni(III) fragment. The spn forms a strong two-electron bond with Ni dz2, in which the bonding orbital primarily is associated with C, corresponding to a strong transfer of charge to C. The antibonding orbital lies above the dx2y2 orbital, unoccupied. Reproduced from Dey et al. [22] with permission of the American Chemical Society, copyright (2007).

reactivity of the proposed intermediates and on the observed stereochemistry of the overall reaction. The presently discussed catalytic mechanisms can in principle be divided into two basic models. In one model the key intermediate features a methylNi(III) species being either formed in a nucleophilic substitution reaction (see Section 4.1) or in an oxidative addition reaction (Section 4.2). In the other model first the thioether sulfur of methyl-coenzyme M binds to the Ni(I), which subsequently results in the release of the methyl group as methyl radical leaving behind a Ni(II)-sulfur bond (Section 4.3). The different models have in common, that they involve directly or indirectly a thiyl radical and a disulfide radical anion as intermediates in the catalytic cycle. Such radicals have been shown to be involved in ribonucleotide reduction [37,38]. The redox potential of the thiyl radical/thiol couple has been estimated to be of the order of +1.3 V [38] relative to the normal hydrogen electrode and that of the disulfide/disulfide radical anion couple to be 1.4 V [39]. These potentials are more positive and more negative than the redox potential of the Ni(III)F430/Ni(II)F430 couple of 4 +1 V and of the Ni(II)F430/Ni(I)F430 couple of –0.6 V, respectively. Met. Ions Life Sci. 2009, 6, 115–132


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4.1. Methylation of Ni(I)F430 in the Active Site via Nucleophilic Substitution It is assumed in this mechanism that the first step in the catalytic cycle is the reaction of the methyl group of methyl-coenzyme M with Ni(I) in a nucleophilic substitution reaction (Figure 4A) yielding methyl-Ni(III) and coenzyme M (reaction 10). Methyl-Ni(III) is then thought to be protonolyzed to methane and Ni(III) in an electrophilic substitution reaction, the proton donor most probably being coenzyme B (reaction 11). Subsequently Ni(III) oxidizes the thiol group of coenzyme M to the thiyl radical (-Sd) (reaction 12) which in turn reacts with coenzyme B to the disulfide radical anion (-Sd-S-)– with a two center-three electron (2c-3e) bond (reaction 13). The latter is a strong reductant capable of re-reducing Ni(II)F430 to Ni(I)F430 (reaction 14) thus closing the catalytic cycle [12]. CH3 -S-CoM þ NiðIÞF430 Ð CH3 -NiðIIIÞF430 þ CoB-S

ð10Þ

CH3 -NiðIIIÞF430 þ CoB-SHÐ CH4 þ NiðIIIÞF430 þ CoB-S

ð11Þ

NiðIIIÞF430 þ CoM-S Ð NiðIIÞF430 þ CoM-Sd

ð12Þ

CoM-Sd þ CoB-S Ð CoM-Sd -S-CoB

ð13Þ

CoM-Sd -S-CoB þ NiðIIÞF430 Ð CoM-S-S-CoB þ NiðIÞF430

ð14Þ

This mechanism is consistent with the finding that MCR-catalyzed ethylcoenzyme M reduction proceeds with inversion of configuration at C1 of its ethyl group [40], since reaction (10) is predicted to proceed with inversion and reaction (11) with retention of configuration. Also the finding that MCR catalyzes the reduction of ethyl-coenzyme M with less than 1% of the catalytic efficiency of methyl-coenzyme M reduction is in agreement with a nucleophilic substitution as first step, which predicts that the electrophilic attack of Ni(I) on C1 of the ethyl group should be sterically hindered [19] (Figure 4A). But there are also problems. The formation of CH3-Ni(III)F430 from CH3S-CoM and Ni(I)F430 has no yet been shown, neither with free Ni(I)F430 nor with Ni(I)F430 bound in MCRred1. Based on DFT calculations, Siegbahn et al. concluded that reaction (10) would be energetically too unfavorable [41,42]. In the case of free Ni(I)F430 this is intuitively understandable since the CH3-S bond (approximately 300 kJ/mol) is much stronger than the CH3-Ni bond (80 kJ/mol) [31]. In agreement with this prediction methyl-coenzyme M is readily formed from MCRMe (generated from Met. Ions Life Sci. 2009, 6, 115–132

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Figure 4. Optimal position of methyl-coenzyme M in the active site of methylcoenzyme M reductase assuming (A) methylation of Ni(I) via nucleophilic substitution [12,15], (B) methylation of the Ni(I) center via oxidative addition [47], and (C) reaction of the Ni(I) center with the sulfur of the thioether bond of methylcoenzyme M [41,42]. The long aliphatic arm of coenzyme B (Figure 1) can reach into the channel only to the extent where its terminal thiol group is in a distance of 8 A˚ from Ni(I).

MCRred1 and methyl bromide; reaction 7) upon addition of coenzyme M [22]. For methyl-coenzyme M to react with free Ni(I)F430 the CH3-S bond in methyl-coenzyme M has to be activated by protonation (or methylation) of the sulfur [1]. But, there is no indication from the crystal structure of MCR that an activation of the CH3-S bond by protonation occurs. The two conserved tyrosines in the active site, which are potential proton donors, are not positioned appropriately [12–15]. It has to be kept in mind, however, that the crystal structure is that of the inactive MCR in the Ni(II) oxidation state and that there is evidence for substantial conformational changes in the active site upon binding of coenzyme B to the active enzyme [17,24]. Methyl-coenzyme M reacts with Ni(I)F430 in MCRred1 only in the presence of coenzyme B, which upon binding to the active enzyme induces a conformational change forcing methyl-coenzyme M and Ni(I) of the prosthetic group to interact. This enforced interaction is deduced from the finding that inactivation of MCRred1 by bromoethanesulfonate and by other suicide inhibitors is dependent on the presence of coenzyme B [19] and that the substrate analogue coenzyme M binds with its thiol group to Ni(I) in MCRred1 only in the presence of coenzyme B as revealed by EPR spectroscopy [43,44]. In single turnover experiments methane formation from methyl-coenzyme M was found to be dependent on coenzyme B [45]. Considering the involvement of methyl-coenzyme M reductase in anaerobic methane oxidation with sulfate, the methane-forming step in the Met. Ions Life Sci. 2009, 6, 115–132


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catalytic cycle must be reversible (reaction 11). Indeed, the reaction of methane, either end-on or side-on, with Ni(III)F430 as described for the activation of C-H bonds by other high-valent metal complexes can be envisaged [46].

4.2. Methylation of Ni(I)F430 in the Active Site via Oxidative Addition This mechanism, which is mainly based on DFT calculations, also involves methyl-nickel bond formation but is otherwise quite different [47]. The cycle begins with the protonation of Ni(I)F430 either on Ni(I) or on the C-ring nitrogen of the corphin ring to yield Ni(I)H1F430 (reaction 15). Ni(I)H1F430 is predicted to oxidatively add CH3-S-CoM (reaction 16) to give a 4-coordinate Ni center (Figure 4B) with the nickel above the plane of the corphin ring, to which the nickel is now coordinated only to two of the four nitrogen atoms of the corphin ring. The two other ligands are the methyl group and SCH2CH2SO2 3 . By binding of CoB-SH, the Ni (and the attached CH3 and SCH2CH2SO2 ligands) moves towards the S-CoB (deprotonated HS3 CoB) cofactor allowing a 2c-3e interaction to form between the two sulfur atoms and allowing the transfer of two electrons from S-CoB to the Ni center. A Ni(II)-coordinated heterodisulfide radical anion is an intermediate. The release of the heterodisulfide yields CH3-Ni(I)H1F430 ¼ CH3-Ni(III)HF430 (reaction 17) from which methane is reductively eliminated (reaction 18) NiðIÞF430 þ CoB-SH Ð NiðIÞHþ F430 þ CoB-S

ð15Þ

NiðIÞHþ F430 þ CH3 -S-CoM Ð ðCH3 -ÞðCoM-S-ÞNiðIIIÞHþ F430

ð16Þ

ðCH3 -ÞðCoM-S-ÞNiðIIIÞHþ F430 þ CoB-S Ð CH3 -NiðIÞHþ F430 þ CoM-S-S-CoB

ð17Þ

CH3 -NiðIÞHþ F430 $ CH3 -NiðIIIÞH F430 Ð CH4 þ NiðIÞF430

ð18Þ

The oxidative addition (reaction 16) and the reductive elimination (reaction 18) both occur via retention of stereochemistry, which appears to exclude this mechanism since, as indicated above, it has been shown that the reduction of ethyl-coenzyme M with coenzyme B proceeds with inversion of stereochemistry at C1 of the ethyl group [40]. However, the catalytic efficiency of MCR with ethyl-coenzyme M is less than 1% of that with methylcoenzyme M [17]. Therefore, it is argued by Duin and McKee [47] that with Met. Ions Life Sci. 2009, 6, 115–132

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such a big difference it is possible that the mechanism followed for the reaction of ethyl-coenzyme M is very different from the one followed for methyl-coenzyme M. In line with this mechanism is the recent finding that MCRred2r is formed from MCRred1 by the coenzyme B induced oxidative addition of the H-S bond of coenzyme M to Ni(I)F430 to give a Ni(III) center, to which a hydride and the sulfur of –SCH2CH2SO2 3 are ligated [24].

4.3. Catalytic Mechanism of Methyl-Coenzyme M Reductase Not Involving Metal-Carbon Bond Formation This mechanism, which is based on predictions of DFT calculations, is generally referred to as the Siegbahn mechanism [41,42]. It starts with the reaction of the thioether sulfur of methyl-coenzyme M with the Ni(I) center in MCRred1 (Figure 4C) yielding CoM-S-Ni(II)F430 and a methyl radical (CHd3) (reaction 19) which subsequently reacts with coenzyme B forming methane and the thiyl radical (-Sd) of coenzyme B (reaction 20). The thiyl radical subsequently reacts with the sulfur attached to Ni(II) resulting in a Ni(II)-coordinated heterodisulfide radical anion (-S-dS-) with a two center three electron (2c-3e) bond, from which the third electron is transferred to the Ni(II) center yielding Ni(I)F430 and CoM-S-S-CoB (reaction 21). CH3 -S-CoM þ NiðIÞF430 Ð CoM-S-NiðIIÞF430 þ CHd3

ð19Þ

CHd3 þ CoB-SH Ð CH4 þ CoB-Sd

ð20Þ

d S-CoB

þ CoM-S-NiðIIÞF430 Ð NiðIÞF430 þ CoM-S-S-CoB

ð21Þ

Reaction (19) is calculated to have a much lower energy barrier than reaction (10), which is the main reason for this mechanistic proposal. In their second paper Siegbahn et al. [41,42] propose that the release of the methyl radical (reaction 19) and its reaction with coenzyme B (reaction 20) proceed in a concerted manner such that the transfer of the methyl group is associated with an inversion of stereo configuration. The mechanism does not explain why ethyl-coenzyme M is so much more slowly reduced to ethane than methyl-coenzyme M to methane and why allyl-coenzyme M is not at all reduced although allyl-coenzyme M is bound in the active site [19]. The argument is that the ethyl radical and the allyl radicals are much better leaving groups than the methyl radical. Steric reasons for why ethyl-coenzyme M and allyl-coenzyme M are reduced much Met. Ions Life Sci. 2009, 6, 115–132


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more slowly or not at all are not evident from the MCR crystal structure [12]. In the active site of MCR there is enough space for the ethyl and the allyl rest to be positioned like the methyl group of methyl-coenzyme M when being reduced by the Ni(I) center (Figure 4C).

5. OBSERVATIONS TO BE FOLLOWED UP Coenzyme M has been shown to bind with its thiol(ate) sulfur to the MCR Ni center in the Ni(III) (MCRox1) state [25,48,49] and Ni(II) (MCRox1-silent) state [12], as well as in the Ni(I) (MCRred2r) state [24,43,44], which might have to be formally described as (H)(CoMS)Ni(III) [23,24]. However, the thioether sulfur of methyl-coenzyme M is expected to show much weaker coordination than the thiol(ate). Recent EPR results on the interaction of methyl-coenzyme M with the Ni(I) center in the MCRred1m state (methyl-coenzyme M bound in the active MCRred1 form) point to an extremely weak coordination with a SNi distance of 43.5 A˚ (unpublished results). Apparently, only the binding of coenzyme B, which seems to induce a conformational change and rearrangement of the coordination sphere, triggers the crucial step, namely the breaking of the CH3-S bond in the substrate. When methyl-coenzyme M is added to MCRred1, the EPR signal changes only slightly. There is also an only minor change when additionally coenzyme B is added despite the fact that now methane is actively formed [16]. The MCRred1 signal is quenched, however, when MCRred1 is supplemented with both methyl-coenzyme M and HS-CoB6 (N-6-thiohexanoyl-O-phospho-Lthreonine) (unpublished results). The coenzyme B homologue has been shown to induce, as coenzyme B, the conformational changes but without being a substrate [17]. The quenching of the signal indicates that methyl-coenzyme M reacted with the Ni(I). A kinetic analysis of the inactivation reaction could reveal which of the considered mechanisms are to be excluded.

ACKNOWLEDGMENTS This work was supported by the Max Planck Gesellschaft, by the Fonds der Chemischen Industrie, and the Schweizerischer Nationalfonds zur Fo¨rderung der Wissenschaftlichen Forschung.

ABBREVIATIONS 2c-3e bond BES

two-center three-electron bond 2-bromoethanesulfonate Met. Ions Life Sci. 2009, 6, 115–132

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BPS BrMe CH3-S-CoM CoM-S-S-CoB CoB-SH CoM-SH CW DFT ENDOR F430 F430M HYSCORE MCR NHE

JAUN and THAUER

3-bromopropane sulfonate methyl bromide methyl-coenzyme M; 2-(methylthio)ethane sulfonate heterodisulfide of coenzyme M and coenzyme B coenzyme B; N-7-thioheptanoyl-O-phosphoL-threonine coenzyme M, 2-thioethanesulfonate continuous wave density function theory electron nuclear double resonance nickel porphin coenzyme F430 pentamethylester of F430 hyperfine sublevel correlation methyl-coenzyme M reductase normal hydrogen electrode

REFERENCES 1. B. Jaun and R. K. Thauer, in Nickel and Its Surprising Impact in Nature, Vol. 2 of Metal Ions in Life Sciences, Ed. A. Sigel, H. Sigel and R. K. O. Sigel, John Wiley & Sons, Ltd, Chichester, UK, 2007, pp. 323–356. 2. S. Shima and R. K. Thauer, Curr. Opin. Microbiol., 2005, 8, 643–648. 3. R. K. Thauer and S. Shima, in Archaea, Evolution, Physiology and Molecuar Biology, Ed. R. Garrett and H.-P. Klenk, Blackwell Publishing, Inc., Malden, USA, 2007, pp. 275–283. 4. R. Thauer and S. Shima, Ann. NY Acad. Sci., 2008, 1125, 158–170. 5. M. Goubeaud, G. Schreiner and R. K. Thauer, Eur. J. Biochem., 1997, 243, 110–114. 6. A. Eschenmoser, Ann. NY Acad. Sci., 1986, 471, 108–129. 7. R. K. Thauer, Microbiology, 1998, 144, 2377–2406. 8. U. Ermler, Dalton Transactions, 2005, 3451–3458. 9. E. C. Duin, in Tetrapyrroles, Ed M. J. Warren and A. G. Smith, Landes Bioscience and Springer Science+Business Media, 2007. 10. R. Piskorski and B. Jaun, J. Am. Chem. Soc., 2003, 125, 13120–13125. 11. B. Jaun, in Properties of Metal Alkyl Derivatives, Vol. 29 of Metal Ions in Biological Systems, Ed. H. Sigel and A. Sigel, Marcel Dekker, New York, 1993, pp. 287–337. 12. U. Ermler, W. Grabarse, S. Shima, M. Goubeaud and R. K. Thauer, Science, 1997, 278, 1457–1462. 13. W. Grabarse, F. Mahlert, S. Shima, R. K. Thauer and U. Ermler, J. Mol. Biol., 2000, 303, 329–344. 14. W. Grabarse, F. Mahlert, E. C. Duin, M. Goubeaud, S. Shima, R. K. Thauer, V. Lamzin and U. Ermler, J. Mol. Biol., 2001, 309, 315–330.

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Met. Ions Life Sci. 2009, 6, 133–150

4 Nickel-Carbon Bonds in Acetyl-Coenzyme A Synthases/Carbon Monoxide Dehydrogenases Paul A. Lindahl Departments of Chemistry and of Biochemistry and Biophysics, Texas A&M University, College Station, TX 77843, USA

ABSTRACT 1. INTRODUCTION 2. REDOX AND CATALYTIC PROPERTIES OF THE A- AND C-CLUSTERS 3. EVIDENCE FOR A Ni-CO BOND IN THE ARED-CO STATE OF THE A-CLUSTER 4. EVIDENCE FOR A Ni-CH3 BOND IN THE METHYLATED INTERMEDIATE OF THE A-CLUSTER 5. EVIDENCE FOR A Ni-C(O)CH3 BOND IN THE ACETYL INTERMEDIATE OF THE A-CLUSTER 6. EVIDENCE FOR A Ni-CO BOND IN THE C-CLUSTER 7. EVIDENCE FOR A Ni-C(O)O-Fe BOND IN THE C-CLUSTER 8. CONCLUSIONS AND FUTURE STUDIES ACKNOWLEDGMENT ABBREVIATIONS AND DEFINITIONS REFERENCES

133 134 137 139 139 140 141 143 144 147 147 147

ABSTRACT: Acetyl-coenzyme A synthases/carbon monoxide dehydrogenases are bifunctional enzymes that catalyze the synthesis of acetyl-CoA and the reversible reduction of CO2 to CO. The active site for the first reaction, called the A-cluster, consists of a [Fe4S4] cubane bridged to a dinuclear nickel subcomponent. The active site for the second reaction, the C-cluster, consists of a [Fe3S4] subsite linked to a Ni-Fe dinuclear Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00133

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site. There is evidence for the formation of five Ni-C bonds, involving methyl, acetyl, carbonyl, and carboxylate groups. In this review, the current evidence for each of these bonds is described. The mechanism of catalysis is discussed, highlighting the role of these species. The unique coordination environments of nickel that may facilitate the formation of organometallic species is discussed. Current puzzles in the field and future research directions that might resolve them are outlined. KEYWORDS: A-cluster C-cluster Fe/S clusters organometallic EPR spectroscopy Mo¨ssbauer spectroscopy infrared spectroscopy

1. INTRODUCTION Acetyl-coenzyme A synthases/carbon monoxide dehydrogenases (ACS/ CODH) are bifunctional Ni-containing enzymes found in certain bacteria and archaea. The organisms that contain such enzymes are anaerobic, mostly thermophilic, and able to grow chemiautotrophically on simple inorganic compounds (e.g., CO2 as a source of carbon). The metabolic and phylogenetic characteristics of the organisms containing this family of enzymes have been reviewed recently [1]. Other recent reviews have focused on the structure and function of the metal centers in these enzymes and on synthetic model complexes [2–4]. The major objective of this review is to describe the evidence for Ni-C bonds contained within these enzymes (when prepared in particular states). There are five possible catalytic intermediates which have been proposed to contain such bonds, involving methyl, acetyl, carbonyl, and carboxylate groups. Evidence for these bonds ranges from unambiguous and definite, to indirect and somewhat speculative. Before describing each candidate organometallic species, and the evidence for it, the structure and physiochemical properties of the active-site metal clusters will be outlined, as will the most popular mechanisms of catalysis. Readers should consult other reviews for a more complete description of the structure of the enzymes and associated metal centers, and for a more detailed comparison of competing proposed catalytic mechanisms. The best-studied ACS/CODH, from the mesophile Moorella thermoacetica, is an 310,000 Da a2b2 tetramer. The two b subunits, which form the central core (Figure 1a), catalyze the reversible oxidation of CO to CO2 (CODH reaction 1). The active site for this activity, called the C-cluster, is CO þ H2 O Ð CO2 þ 2e þ 2Hþ

ð1Þ

a NiFe4S4 cluster (Figure 1b) (for complete structural details, see [5–10]). The C-cluster consists of an [Fe3S4] subsite linked to a [NiFea] subsite. Two of the three m2-bridging sulfides of the [Fe3S4] subsite additionally coordinate to the Ni ion while the third m2-bridging sulfide additionally coordinates Met. Ions Life Sci. 2009, 6, 133–150

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Figure 1. Structures of ACS/CODH and associated metal centers. (a) symbolic protein structure of ACS/CODH. Each a subunit contains three domains which are in slightly different conformations. Each a subunit contains a single metal center called the A-cluster (green). The two b subunits form the central core of the enzyme. These subunits contain three types of metal centers, including the active-site Cclusters (green) and electron-transfer clusters B and D (brown); (b) structure of the Ccluster; (c) structure of the A-cluster.

to a unique Fe called Fea. The Ni and Fea can be bridged by a hydroxo group (almost certainly derived from the substrate H2O) [10], or by sulfido [5,9] and perhaps other anions such as CN [11], SCN, OCN and N 3 groups [12,13]. The Ni Fea bridging site can also be unoccupied [7,8]. The Ccluster Ni is also coordinated to a cysteine thiolate, resulting in a planar three-coordinate T-shaped geometry when the bridging position is empty Met. Ions Life Sci. 2009, 6, 133–150

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and a square-planar geometry when it is occupied. Completing the coordination sphere of Fea is a cysteine thiolate and a histidine imidazole. Three cysteine thiolates affix the [Fe3S4] subsite to the protein. The C-cluster is buried ca. 18 A˚ below the protein surface [7]. Two [Fe4S4] clusters in the b subunits form a ‘‘wire’’ that electronically connects the C-cluster to redox agents in the solvent. The B-cluster is proximal to the C-cluster and located 11 A˚ away from the C-cluster (Figure 1a). The distal D-cluster, about 10 A˚ away from the B-cluster, bridges the two b subunits and is essentially at the protein surface. The A-cluster active site within the a subunits (Figure 1a) catalyzes the acetyl-CoA synthase (ACS) activity (reaction 2). CH3 -Co3þ FeSP þ CO þ CoASH Ð CH3 -CðOÞ-CoA þ Co1þ FeSP þ Hþ

ð2Þ

CoFeSP, the corrinoid-iron-sulfur protein, is the specific methyl group donor to ACS/CODH. In a separate reaction, CoFeSP accepts a methyl group from methyl-tetrahydrofolate (H4F), in accordance with reaction (3), which is catalyzed by a methyltransferase. CH3 -H4 F þ Co1þ FeSP þ Hþ Ð CH3 -Co3þ FeSP þ H-H4 F

ð3Þ

The A-cluster consists of a [Fe4S4] cubane bridged through a cysteine residue to a Ni ion (called proximal Nip) which is additionally bridged, through 2 cysteine residues, to a second Ni ion called distal Nid (Figure 1c) (for complete structural details, see [7–8,14]). Nip is square-planar, including the three m2-bridging cysteine thiolates and an endogenous unidentified ligand which is presumably replaced by substrates during catalysis. The N2S2 square-planar geometry of Nid is completed by the coordination to two amide nitrogens originating from the protein backbone. The acetyl-CoA decarbonylase/synthases (ACDS) represent another class of enzymes within the Ni-containing CODH family [15]. These trifunctional abgde enzymes have ACS, CODH and methyltransferase activities, and they function in acetoclastic methanogenesis. The isolated a subunit of ACS/ CODH and the homologous subunit from other enzymes can be prepared by recombinant genetic methods, and are active catalysts after incubation in NiCl2 [14–18]. The monomeric ACS from Carboxydothermus hydrogenoformans has been structurally characterized [14]. There are also monofunctional Ni-dependent enzymes that catalyze only the CODH reaction, with the b2 dimeric enzyme from Rhodospirillum rubrum (CODHRr) being the best-studied [19]. Met. Ions Life Sci. 2009, 6, 133–150


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2. REDOX AND CATALYTIC PROPERTIES OF THE A- AND C-CLUSTERS The C-cluster can be stabilized in 4 redox states, including Cox (S ¼ 0), Cred1 (S ¼ 1/2, with EPR principal g-values of 2.01, 1.81, 1.65 for ACS/CODH), Cint (S ¼ 0 or integer), and Cred2 (S ¼ 1/2, with EPR principal g-values of 1.97, 1.87, 1.75, again for ACS/CODH) [11,20,21]. The Cox state is most oxidized and the others are progressively more reduced in one-electron increments. Reasonable electronic configurations for the metal ions in the Cox and Cred1 states include {[Fe31, Fe21, Fe21] [Ni21 Fea31]} and {[Fe31, Fe21, Fe21] [Ni21 Fea21]}, respectively [11,22], while those for Cint and Cred2 are less certain. The best-supported or ‘‘standard’’ CODH catalytic mechanism is shown in Figure 2a. The C-cluster becomes activated for catalysis when it is reduced from the Cox to the Cred1 state [23]. A hydroxide ion bridges the Ni and Fea in the Cred1 state [13,24]; this ion is derived from the substrate H2O and it serves as nucleophile to attack a Ni-bound carbonyl carbon (see below) [25]. Deprotonation of the resulting carboxylic acid leads to a ½Ni-CO2 -Fea intermediate [10]. Subsequent decarboxylation leads to the Cred2 state. Two one-electron oxidations, the binding of water, and the deprotonation of that water return the C-cluster to the Cred1 state, completing the catalytic cycle. The A-cluster can be stabilized in three well-established states (Aox, CH3Aox, and Ared-CO) and perhaps in two less-established states (Ared-act and CH3C(O)-Aox). The S ¼ 0 Aox state is obtained when ACS/CODH (or the isolated Ni-activated a subunit) is prepared in an inert-atmosphere glove box without added reductants or oxidants. The electronic configuration of 2þ Aox appears to be {[Fe4S4]21 Ni2þ p Nid } [26,27]. The S ¼ 1/2 Ared-CO state is obtained by treating ACS/CODH with CO or by treating the a subunit with CO and a low-potential reductant like dithionite or Ti(III) citrate. The Ni becomes reduced and bound with CO, affording the electronic configuration 2þ {[Fe4S4]21 Ni1þ p -CO Nid }. The Ared-CO state exhibits the well-studied NiFeC EPR signal [28]. In the so-called diamagnetic mechanism (Figure 2b), Ared-CO represents an inhibited form of catalysis. However, in the paramagnetic mechanism, Ared-CO is considered to be a catalytic intermediate [2,3,29,30]. According to the diamagnetic mechanism, the A-cluster in the Aox state is reduced by two low-potential electrons to the reductively-activated Ared-act state before it is active for catalysis [27]. The electronic configuration for this state is controversial, but it is probably either {[Fe4S4]21 Ni0p Ni2þ d } or 2þ Ni } [31–34]. The diamagnetic CH -A state is the {[Fe4S4]11 Ni1þ 3 ox p d methylated intermediate (described below) with electronic configuration 2þ {[Fe4S4]21 Ni2þ p -CH3 Nid } [27]. This state is obtained by reducing ACS/ CODH (or the isolated a subunit) with a low-potential reductant, followed Met. Ions Life Sci. 2009, 6, 133–150

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Figure 2. The ‘‘standard’’ CODH mechanism (a) and the diamagnetic ACS mechanism (b).

by a methyl group transfer reaction with CH3-Co31FeSP [35–40]. The CH3C(O)-Aox state represents the acetylated intermediate, obtained upon CO binding and insertion. The acetyl group is transferred from the A-cluster to CoA, forming the product acetyl-CoA and regenerating the Ared-act state for another round of catalysis. The detailed kinetics associated with this mechanism have been mathematically modeled [41]. Met. Ions Life Sci. 2009, 6, 133–150


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3. EVIDENCE FOR A Ni-CO BOND IN THE ARED-CO STATE OF THE A-CLUSTER Evidence for this Ni-C bond comes from EPR [28], IR [18,42], and ENDOR [43] spectroscopies, as well as from computational studies [33] and synthetic analogs [44,45]. The S ¼ 1/2 Ared-CO state exhibits the axial NiFeC EPR signal, with gperp ¼ 2.074 and gpar ¼ 2.028 [28]. The signal broadens due to 13 C hyperfine interactions when ACS/CODH (or a) is exposed to 13CO. This broadening indicates that the nuclear spin of 13CO (I ¼ 1/2) is coupled to the S ¼ 1/2 electronic spin of the A-cluster. Ragsdale and coworkers have correlated this binding to an intense IR peak at 1996 cm1 [18,42]. An IR peak at this energy represents a terminal (end-on) CO bound to a metal ion. The evidence that this CO binds to Nip rather than to Nid or to the [Fe4S4] component of the A-cluster is indirect. Mo¨ssbauer and UV-vis spectroscopies indicate that in the Ared-CO state, the [Fe4S4] cubane is in the 2+ core oxidation state [26]. The binding of CO to an [Fe4S4]21 cubane has not been reported, and so this seems unlikely. Although the oxidation state of Nip cannot be observed directly by Mo¨ssbauer, the fact that the system is S ¼ 1/2 suggests that either Nip or Nid (but not both) is in the 1+ oxidation state. The binding of CO to synthetic Ni11 model compounds that mimic the ACS site suggests that CO binds to Nip11 in the Ared-CO state [44,45]. The presence of hyperfine broadening when 61Ni is incorporated into the proximal site of the A-cluster and the lack of such broadening when 61Ni is 2þ incorporated into the distal site [31,46] also favors the fNi1þ p Nid g configuration. Further support for this assignment are Ni model complexes with N2S2 square-planar geometries and amide nitrogens for which Nid-like Ni ions can readily become oxidized to the Ni31 state, but cannot be reduced to the 1+ state [47].

4. EVIDENCE FOR A Ni-CH3 BOND IN THE METHYLATED INTERMEDIATE OF THE A-CLUSTER Evidence for this state is substantial but indirect. Using 14C-labeled methyl groups, Pezacka and Wood discovered in the late 1980’s that a methyl group from the CoFeSP could be transferred to reduced ACS/CODH [35]. The resulting methylated adduct is stable and functional, as subsequent addition of CO and CoA affords acetyl-CoA [38]. Ragsdale and coworkers quantified the requirement for reducing the enzyme prior to methyl group transfer [36] and established that the methyl group is transferred by a nucleophilic displacement mechanism [37]. Detailed stopped-flow kinetics of ACS/CODH and a afforded forward and reverse rate constants as well as an equilibrium Met. Ions Life Sci. 2009, 6, 133–150

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constant for the process [38,39,41]. In conflict with the original paramagnetic mechanism [48], the methylated intermediate was found to be EPR-silent [38]. Mo¨ssbauer studies showed that the cubane in this state was in the oxidized 2+ core oxidation state and apparently diamagnetic [26,27]. The site on the A-cluster that is methylated has not been identified unambiguously. However, there is circumstantial evidence that this site is Nip [38]. Nip can be removed by treating the enzyme with 1,10-phenanthroline (phen) [46,49]. This abolishes ACS catalytic activity and the ability of ACS/CODH to accept a methyl group from CH3-Co31FeSP [38,39]. The reaction is specific for Nip and is reversible. Thus, if phen-treated enzyme is incubated in NiCl2, the Ni is installed into the A-cluster and catalytic activity and methyl group transfer ability return. The simplest explanation of why the methyl group cannot bind phen-treated enzyme is that it binds Nip which is absent in this form of the enzyme. In further support of this explanation, the removal of Nip by phen is inhibited when the enzyme is methylated [36], suggesting that phen cannot coordinate Nip when the methyl group is bound to it. Calculations suggest that the methylated intermediate might have a square-planar geometry [32]. The bidentate ligand phen, which probably coordinates Nip at two cis-adjacent sites, might not be able to bind and remove methyl-bound Nip because the methyl occupies one of these sites.

5. EVIDENCE FOR A Ni-C(O)CH3 BOND IN THE ACETYL INTERMEDIATE OF THE A-CLUSTER The evidence for an acetylated state of ACS/CODH is indirect but compelling nonetheless. Since 1986, the enzyme has been known to undergo the acetyl-CoA/*CoA exchange reaction [50–52] which implies an acetyl-bound intermediate, e.g., as illustrated in reactions (4) and (5). CH3 -CðOÞ-CoA þ E Ð CH3 -CðOÞ-E þ CoA

ð4Þ

CH3 -CðOÞ-E þ CoA Ð CH3 -CðOÞ- CoA þ E

ð5Þ

In these reactions, E represents the reduced form of either ACS/CODH or an ACDS homolog and *CoA is either radio-labeled CoA or dephosphoCoA. Grahame and coworkers analyzed the steady-state kinetic behavior of this type of exchange reaction to changes in the concentrations of substrates and products [53]. Their results clearly show that exchange reaction proceeds by a ping-pong mechanism, as suggested in reactions (4) and (5), implying an Met. Ions Life Sci. 2009, 6, 133–150


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acetyl-bound intermediate. They calculate that hydrolysis of the acetylbound enzyme intermediate (reaction 6), CH3 -CðOÞ-E þ H2 O Ð CH3 -COOH þ E þ Hþ

ð6Þ

proceeds spontaneously with DG00 E 9.5 kcal/mol [53]. This indicates a ‘‘high energy’’ bond, consistent with the acetyl group being bound to a metal center. The acetyl-bound intermediate hydrolyzes slowly (in B30 min) at room temperature [38], indicating that it is isolatable and susceptible to spectroscopic analysis. The acetylated species is EPR-silent [38] and exhibits Mo¨ssbauer spectra indistinguishable from that of methylated enzyme (cubane in the diamagnetic 2+ core oxidation state) [27]. Both results suggest that the acetylated intermediate is diamagnetic. Strictly speaking, the site on ACS/CODH where the acetyl group binds is unknown but Nip is perhaps the only reasonable possibility. Given the evidence that the methyl group binds to Nip (see above), it is simplest to assume that the corresponding acetyl group would be bound at the same metal ion. Synthetic diamagnetic Ni21-C(O)CH3 complexes are known [44], which supports this possibility. Considered collectively, the acetylated intermediate would appear 2þ to have the electronic configuration f½Fe4 S4 2þ Ni2þ p -CðOÞCH3 Nid g.

6. EVIDENCE FOR A Ni-CO BOND IN THE C-CLUSTER The case for this state is perhaps the weakest of all those mentioned in this review, but the evidence for it, viewed collectively, is still substantial. The best evidence comes from IR spectra of CO-treated ACS/CODH which exhibit peaks at 2078, 2044, 1970, 1959, and 1901 cm1 [42]. These peaks have been assigned to C-cluster species bound with terminal COs. These peaks disappear with time after CO exposure, consistent with the oxidation of the associated carbonyl species to CO2 as the enzyme slowly (over the course of B100 min) oxidizes. Slightly different forms of the C-cluster exist in solutions of ACS/CODH, some of which are functional and others of which are not [11,54]. Similarly, there is structural evidence for slightly different locations of some atoms in C-cluster structures [8–10]. The five IR peaks might arise from this heterogeneity, each reflecting a single CO bound at multiple C-cluster structural variants [42]. An alternative interpretation, which we find less appealing, is that multiple CO molecules bind to different sites on a single functional C-cluster. A modicum of evidence for a Ni-CO bond at the C-cluster comes from the reported 2.8 A˚ resolution X-ray diffraction structure reported for CO-treated CODH from Rhodospirillum rubrum [6]. Electron density indicated an Met. Ions Life Sci. 2009, 6, 133–150

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exogenous ligand on the Ni of the C-cluster, and this density was modeled as CO. The C-cluster in the 1.9 A˚ resolution structure of CO-treated ACS/ CODH also contained electron density that indicated an exogenous ligand coordinated to the axial position of the Ni, and this density was also modeled as CO [8]. The C-cluster structure from the non-CO treated CODH from Carboxydothermus hydrogenoformans did not indicate such coordination [5], but a subsequently reported structure of a CO-treated sample also exhibited electron density at the Ni, which was again modeled as CO [9]. Collectively, these X-ray diffraction studies suggest that CO binds to the axial site of the Ni in the C-cluster when the enzyme is exposed to CO. Difficulties in modeling the electron density due to this species may arise from structural heterogeneity of the center. Interestingly, there is a structural correlation between the absence of the bridging sulfide (or hydroxide) and exposure to CO [9]. CO-treated ACS/ CODH exhibits the Cred2 EPR signal, which suggests that CO binds the Ccluster in the Cred2 state. ENDOR indicates a strongly coupled proton (which probably reflects the bridging OH) that is associated with the Cred1 state but not with Cred2 [55]. The absence of bridging OH or SH species in structures where CO is bound to the Ni of the C-cluster is consistent with CO binding stably to the Cred2 state. CO probably also binds to the Cred1 state, but in this case the bound CO probably reacts immediately with the bridging OH group to afford the CO2-bound intermediate described below. One conundrum associated with this hypothesis is that the Cred2 EPR signal shows no evidence of CO binding in CO-treated samples; in fact, the same nominal signal can be obtained by treating the enzyme with the reductant dithionite [20,21]. One explanation considers that the local geometry about the Ni ion in the C-cluster is planar such that the Ni is diamagnetic Ni21. ENDOR and Mo¨ssbauer studies have concluded that the Ni of the C-cluster is weakly coupled (if at all) to the unpaired electron giving rise to the Cred1 EPR signal [11,55]. Similarly, substitution of C-cluster Ni of CODHRr with diamagnetic Zn21 gave rise to a similar EPR signal, suggesting that the EPR signal derives mainly from the Fe/S portion of the Ccluster [56]. Thus, the binding of CO to the Ni may not noticeably alter the Cred2 EPR signal. A similar situation may hold for the CO2-bound form of the C-cluster (see below). The binding of CO to synthetic Ni21 complexes is not common but is established for particular coordination environments [57–59]. Cyanide is a tight slow-binding inhibitor of CODH that was originally proposed to bind exclusively to Fea, due to the observation of a CNdependent shift in the Fea Mo¨ssbauer parameters [11]. However, the bridging of sulfide between Fea and the Ni of the C-cluster [9] and the demonstration that sulfide also inhibits CODH activity suggests that CN (and other anion inhibitors including cyanate, thiocyanate, and azide [12]) might Met. Ions Life Sci. 2009, 6, 133–150


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also bridge the Ni and Fea [13]. Subsequent exposure to CO reverses the inhibition of cyanide and sulfide, such that the activity of CODH recovers slowly (B1 hr) in 1 atm CO, even in the presence of these anions. Anderson and Lindahl originally suggested that CO did this by binding to a ‘‘modulator site’’ [60], but this hypothetical species was eventually assigned to the Ni itself [61]. Activation in this context probably corresponds to the dissociation of these anions and to the binding of the substrate hydroxide ion at the bridging position. The most intriguing aspect of this is that the binding of CO (probably to the Ni of the C-cluster in the Cred1 state) promotes the dissociation of the bridging species, be it an anion inhibitor or the substrate hydroxide. This suggests a sequence of electronic effects initiated by the binding of CO to Ni, which causes the Ni-OH bond to weaken or break [13]. This would free a lone pair of electrons on the hydroxide ion, enhancing the nucleophilic properties of the hydroxide ion which would now be bound solely to Fea. This would prompt the hydroxide to attack the Ni-bound carbonyl carbon, generating a Ni-bound carboxylic acid. Deprotonation of this species would result in the CO2-bound species observed recently by Xray diffraction (see below). Thus, there appears to be a mechanistic domino effect at work here, initiated by the binding of CO to the Ni of the C-cluster in the Cred1 state and terminating in the CO2-bound form of the C-cluster.

7. EVIDENCE FOR A Ni-C(O)O-Fe BOND IN THE C-CLUSTER The evidence for the CO2-bound intermediate state is strong and unambiguous, as this state has been observed recently in an X-ray diffraction structure reported by Jeoung and Dobbek [10]. To obtain their structure, they reduced the sample to the Cred2 state and then added bicarbonate. The resulting structure shows CO2 bound in Z1 fashion to the Ni through the C, with a Ni-C bond distance of 1.96 A˚. One O of the CO2 is coordinated to Fea and is hydrogen-bonded to a conserved lysine; the other O is not coordinated to either metal ion but is hydrogen-bonded to a conserved histidine. Demonstration of this state has challenged our understanding and previous interpretations of reported spectroscopic, redox, and catalytic properties of the C-cluster. Assuming the standard mechanism described above, the C-cluster with CO2 bound should be in the Cred1 redox state (if one assigns the two electrons of the presumed Ni-C bond to the C). But shouldn’t the EPR signal of the CO2-bound C-cluster in this state differ from that obtained in the absence of CO2? Indeed, CO2 exposure slightly shifts the Cred1 EPR g-values [21,62]. However, this would naively imply that CO2 binds the C-cluster in the Cred1 state (whereas the standard model assumes Met. Ions Life Sci. 2009, 6, 133–150

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that CO2 binds in the Cred2 state, and abstracts two electrons from the cluster which are used to form the Ni-C bond). Other evidence would seem to support this idea. Exposing ACS/CODH to the CO2-analog and competitive inhibitor CS2 [63] (in a reducing environment) causes the Cred1 EPR signal to disappear and a novel signal to slowly form [61]. Exposing particular batches of CODH, which are unable to be reduced to the Cred2 state, to CO2/ dithionite can ‘‘cure’’ such batches, allowing them to attain the Cred2 state. How could CO2 have this effect unless it could bind to the enzyme when the C-cluster is in a state more oxidized than Cred2? Also, CO2 (in the presence of dithionite) causes CN to dissociate from the C-cluster, mimicking the effect of CO (which presumably originates from the binding of CO to the Cred1 state). A related puzzle is that CO2 does not shift the Cred2 EPR signal, as would be expected if CO2 bound to the Cred2 state. On the other hand, the reduction potential of Cred1/Cred2 increases in the presence of CO2 [62], as would be expected if CO2 bound to the Cred2 state. Clearly, more studies are required to clarify these issues. A related complication is highlighted by the similar T-shaped coordination geometries of the Ni ion of the C-cluster in both Cred1 and Cred2 states [10]. This seems incongruent with the possibility that the two electrons used to reduce Cred1 to Cred2 locate on the Ni ion, as the reduction of a planar Ni21 ion to a Ni0 species would tend to shift the geometry to tetrahedral. Rather, the structural invariance of the Ni between these two states suggests that the two electrons used to reduce Cred1 to Cred2 are delivered to the [Fe3S4] subsite of the cluster (Fea is already ferrous in the Cred1 state and would probably not accept another electron), and that CO binds to Ni21. However, if the proposed electronic configuration for Cred1 is correct, it seems unlikely that two additional electrons could be accommodated by the [Fe3S4] subsite. A third possibility is that in the Cred2 state, one electron localizes on the Ni (forming Ni11) and the second electron localizes on the [Fe3S4] subsite. A Ni11 designation is favored by mechanistic and computational studies of synthetic Ni complexes that bind CO2 and reduce it to CO [64–66]. Clearly, further studies are required to understand better the details of the Ni oxidation states in the various C-cluster redox states.

8. CONCLUSIONS AND FUTURE STUDIES ACS/CODH and the other members of the Ni-dependent family of CODHs are rare examples in biology where M-C bonds form and where the steps of the catalytic mechanism are characteristic of well-studied organometallic reactions. In this review, we have discussed the evidence for five such bonds in this enzyme, including Nip-CO, Nip-CH3, and Nip-C(O)CH3 Met. Ions Life Sci. 2009, 6, 133–150


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intermediates of the A-cluster and Ni-CO and Ni-CO2-Fe bound intermediates of the C-cluster. When the Co-C bond of the methylcolabamin CH3-Co31FeSP is included, it would appear that ACS/CODH/CoFeSP system holds the ‘‘world record’’ in biology in terms of numbers of Ni-C bonds and organometallic mechanistic steps. What properties of these enzymes and the organisms that contain them facilitate these organometallic species and reaction steps? Clearly, an anaerobic environment is required to stabilize these bonds and preclude reactions of O2 with metal-bound thiolates [67]. The involvement of nickel must also be important, perhaps specifically when coordinated to bridging thiolates and sulfide ions. We have been struck by the similar coordination environments of Nip of the A-cluster and the Ni of the C-cluster (Figure 3); both have three-coordinate T-shaped planar geometries with three sulfur ligands, all but one of which is bridged by other metal centers. This bridging arrangement may neutralize the negative charges on the sulfurs, allowing the coordinated Ni to become readily reduced (either to the Ni11 or Ni0 state) and then serve as a nucleophile to attack a methyl group (in the case of the A-cluster) or CO2 (in case of the C-cluster). The square-planar geometries that apparently result from these attacks suggest that these substrates are bound to diamagnetic Ni21 ions.

Figure 3. Comparison of the Nip and C-cluster Ni coordination environment and reactivity. See text for details. Met. Ions Life Sci. 2009, 6, 133–150

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Where should we go from here? Clearly, obtaining high-quality X-ray diffraction structures of the remaining four Ni-C intermediates will be critical in providing conclusive evidence for their existence. With this structural information in hand, additional computational studies will be critical in defining the ACS and CODH catalytic mechanisms at the atomic level of detail. With regard to the ACS mechanism, the most critical issue (in my mind) is whether the Ared-act state represents a {[Fe4S4]21 Ni0p } or a {[Fe4S4]11 Ni1þ p } configuration. The order of substrate addition must also be settled; recent studies suggest a random mechanism of substrate addition, with either CO or the methyl group adding first and second, and CoA adding last [29]. The enzyme contains an exotic tunnel that connects the Cand A-clusters [7,8]. This tunnel serves not simply to channel CO from the site where it is made (the C-cluster) to the site where it is consumed (the Acluster), but probably to control delivery of CO to the A-cluster such that it arrives at the appropriate step of catalysis [68,69]. How this view of the tunnel’s function can be reconciled with the evidence for the random addition of CO and methyl groups will require further mechanistic studies. With regard to the CODH mechanism, perhaps the most pressing problem is the electronic configuration of Cred2 and a better understanding of the effects of CO/CO2 on the EPR spectra of the C-cluster. Mo¨ssbauer spectroscopy could be used to evaluate iron oxidation states and spin-coupling arrangements, but the problem is that the C-cluster represents at best B30% of the iron in ACS/CODH. Given the heterogeneity of the metal centers in the enzyme, it would appear that only B8% of the Fe in a sample of ACS/ CODH actually contributes to the Cred2 state. Moreover, the B-cluster is reduced and magnetic whenever the C-cluster is in the Cred2 state, and the magnetic spectral features of the two centers overlap. Mutant strains should be developed in which the A-, B-, and D-clusters are either abolished or locked into diamagnetic states, as this would make the study of the electronic properties of the Cred2 state feasible. Another interesting research path would be to probe the CO2-bound state of the C-cluster using 13C and magnetic resonance techniques. If the C-cluster when bound with CO2(CO) is in the Cred1(Cred2) state, perhaps even weak hyperfine interactions could be observed using ESEEM spectroscopy. Ultimately, the study of these enzymes might facilitate the more practical goal of designing new metalloenzymes that catalyze other desirable organometallic reactions. Given the plethora of Ni-C bonds in ACS/CODH, it seems that this enzyme would be an ideal platform upon which such studies could ensue. We have recently attempted (perhaps successfully) to incorporate Pd and Pt into the proximal site of the A-cluster [70]. Future combinations of transition metal coordination chemistry with recombinant genetic engineering might prove successful in developing new bio-organometallic catalysts. Met. Ions Life Sci. 2009, 6, 133–150


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ACKNOWLEDGMENT I would like to thank the National Institutes of Health (GM46441) graciously funding my laboratory to work on this enzyme for the past 15 years.

ABBREVIATIONS AND DEFINITIONS ACDS ACS ACS/CODH Aox Ared-act Ared-CO CH3-Aox CH3C(O)-Aox CoA CODH CODHRr CoFeSP Cox Cred1, Cint, Cred2 ENDOR EPR ESEEM H4F IR Nid Nip phen UV

acetyl-CoA decarbonylase/synthase acetyl-coenzyme A synthase the bifunctional enzyme from Moorella thermoacetica fully oxidized state of the A-cluster the reductively activated state of the A-cluster form of the A-cluster one electron more reduced than Aox and bound with CO the methylated state of the A-cluster the acetylated state of the A-cluster coenzyme A carbon monoxide dehydrogenase CODH from Rhodospirillum rubrum corrinoid-iron-sulfur protein fully oxidized state of the C-cluster redox states of the C-cluster that are 1, 2, and 3 electrons more reduced, respectively, relative to Cox electron nuclear double resonance electron paramagnetic resonance electron spin echo envelop modulation tetrahydrofolate infrared the distal Ni of the A-cluster the proximal Ni of the A-cluster 1,10-phenanthroline ultraviolet

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5 Structure and Function of [NiFe]-Hydrogenases Juan C. Fontecilla-Camps Laboratoire de Cristallographie et de Cristallogene`se des Proteines, Institut de Biologie Structurale J. P. Ebel (CEA-CNRS-UJF), 41 rue Jules Horowitz, F-38027 Grenoble Ce´dex 1, France <[email protected]>

ABSTRACT 1. INTRODUCTION 2. HYDROGENASE STRUCTURE 2.1. The Three-Dimensional Folding 2.2. The Active Site at Medium Resolution 2.3. The Active Site at Higher Resolution 3. HYDROGENASE MATURATION AND ACTIVE SITE ASSEMBLY 3.1. Maturation of the Large Subunit 3.2. Cyanide, Carbon Monoxide, and Iron Insertion 3.3. Nickel Insertion 3.4. Proteolytic Cleavage of the Large Subunit C-Terminal Extension 4. ELECTRON TRANSFER 5. PROTON TRANSFER 6. OXIDIZED INACTIVE STATES OF THE [NiFe]HYDROGENASE ACTIVE SITE 7. SUBSTRATE BINDING AND CATALYSIS 8. CONCLUDING REMARKS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00151

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ABSTRACT: [NiFe(Se)]-hydrogenases are hetero-dimeric enzymes present in many microorganisms where they catalyze the oxidation of molecular hydrogen or the reduction of protons. Like the other two types of hydrogen-metabolizing enzymes, the [FeFe]- and [Fe]-hydrogenases, [NiFe]-hydrogenases have a Fe(CO)x unit in their active sites that is most likely involved in hydride binding. Because of their complexity, hydrogenases require a maturation machinery that involves several gene products. They include nickel and iron transport, synthesis of CN (and maybe CO), formation and insertion of a FeCO(CN)2 unit in the apo form, insertion of nickel and proteolytic cleavage of a C-terminal stretch, a step that ends the maturation process. Because the active site is buried in the structure, electron and proton transfer are required between this site and the molecular surface. The former is mediated by either three or one Fe/S cluster(s) depending on the enzyme. When exposed to oxidizing conditions, such as the presence of O2, [NiFe]-hydrogenases are inactivated. Depending on the redox state of the enzyme, exposure to oxygen results in either a partially reduced oxo species probably a (hydro)peroxo ligand between nickel and iron or a more reduced OH– ligand instead. Under some conditions the thiolates that coordinate the NiFe center can be modified to sulfenates. Understanding this process is of biotechnological interest for H2 production by photosynthetic organisms. KEYWORDS: active site assembly electron transfer enzyme maturation hydrogen oxidation iron with CO and CN coordination [NiFe]-hydrogenases proton transfer

1. INTRODUCTION Hydrogen is utilized by microorganisms as a source of reducing power or produced when protons are used as final electron acceptors. The reaction is mediated by metallo-enzymes called hydrogenases. The need of catalytic transition metal centers is explained by the significant increase in the acidity of molecular hydrogen when bound to metal [1]. Three phylogenetically unrelated classes of these enzymes exist: [NiFe]and [FeFe]-hydrogenases with hydrogen being the only substrate or product, according to the following reaction: H2 Ð 2 Hþ þ 2e

ð1Þ

In the third class, hydrogen uptake is coupled to methenyltetrahydromethanopterin reduction [2]. Recently, the latter has been called [Fe]hydrogenase because, as opposed to the other two enzymes, its active site contains a single metal ion. Previously, the structures of the apo-enzyme and a photolyzed derivative of its Fe-binding cofactor were reported [2]. There is now a crystal structure available for the holoenzyme (S. Shima, personal comm. [97]). There are five structures of [NiFe]-hydrogenases available, all from sulfate-reducing bacteria [3–7]. For [FeFe]-hydrogenase, there are structures from the sulfate-reducing bacterium Desulfovibrio desulfuricans ATCC 7757 [8] and from Clostridium pasteurianum [9]. The Met. Ions Life Sci. 2009, 6, 151–178

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former is a hydrogen-oxidizing periplasmic enzyme whereas the latter is cytoplasmic and produces hydrogen. A low-spin Fe(CO)x motif is common to the three enzyme classes indicating that it represents Nature’s solution to hydrogen catalysis. The active sites of [NiFe]- and [FeFe]-hydrogenases are buried within the structure, requiring electron and proton transport between the catalytic center and the molecular surface. Also, hydrogen has to access the active site or diffuse from it depending on the direction of reaction (1). Because of the unusual structure of the active sites of these enzymes, a considerable effort has been invested in understanding their assembly and significant progress has resulted from this. Last but not least, a recent and quite popular subject of study concerning these enzymes is their degree of sensibility to molecular oxygen. Besides being of basic scientific interest, this study is important when hydrogenases are considered as catalysts in bio-fuel cells or in the industrial production of bio-hydrogen. Prior to the structural analyses of hydrogenases, a series of electron paramagnetic resonance (EPR) spectroscopic studies had shown that the oxidized enzyme is characterized by two paramagnetic states, called Ni-A and Ni-B that disappeared upon activation with hydrogen or other reducing agents. Because enzyme in the Ni-A state was difficult to activate, it was also called the unready form. On the other hand, the Ni-B species was rapidly activated and was consequently considered to be in a ready state. The remaining EPR-active species, obtained under reducing conditions, was called Ni-C and was considered to be part of the catalytic cycle. Two diamagnetic species were observed upon reduction of the Ni-B and Ni-C states and were called Ni-SI and Ni-R, respectively (Figure 1). Theoretical fitting of D. gigas [NiFe]-hydrogenase redox titrations favored the Two Electron Difference (TED) model with the active Ni-B form being two electrons more oxidized than the reduced ready Ni-C state. Thus, the sequence Ni-A/B-NiSI-Ni-C-Ni-R was postulated to represent one-electron reducing steps.

2. HYDROGENASE STRUCTURE 2.1. The Three-Dimensional Folding Crystals of Desulfovibrio (D.) vulgaris Miyazaki [NiFe]-hydrogenase [4] and the D. gigas enzyme [10] were first reported in 1987. However, the first structure was not published until 1995, from a better quality crystal form of the latter enzyme [3]. The crystals were grown by the vapor diffusion method and diffraction data were collected in house with a rotating anode Cu-Ka Xray source at room temperature. X-ray data had to be collected immediately Met. Ions Life Sci. 2009, 6, 151–178

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Figure 1. Ni EPR spectra of the unready Ni-A, ready Ni-B, and active Ni-C paramagnetic species. The spectra of the diamagnetic Ni-SI and Ni-R are also depicted. The successive Ni-B/SI/C/R are thought to be separated by one-electron reductions [84]. Adapted from [92].

after crystal growth because the crystals became colorless and rapidly lost their diffracting power upon exposure to air. A complete 2.85 A˚ resolution X-ray data set was collected from several native crystals and many others were tested for heavy atom derivatives in soaking experiments. The crystal structure was subsequently solved by a combination of multiple isomorphous replacement (MIR) and non-crystallographic symmetry density averaging over the two enzyme heterodimers in the asymmetric unit. The two subunits establish an extensive contact interface of about 3,500 A˚2 (Figure 2). The small subunit has a N-terminal flavodoxin-like Met. Ions Life Sci. 2009, 6, 151–178


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Figure 2. Stereo pair of the crystal structure of [NiFe]-hydrogenase from Desulfovibrio gigas. The large and small subunits are depicted in red and blue, respectively. The three Fe/S clusters and the active site are represented by spheres. Color codes: yellow for S, red for Fe, and green for Ni. Figures 2, 3, 5, 7 and 8 were prepared with the VMD program [96].

domain that represents 65% of the whole polypeptide chain. Superposition of this domain to 89 Ca atoms of a clostridian flavodoxin [11] results in a root mean square deviation of about 2.7 A˚ [3]. The domain coordinates a [Fe4S4] cluster, topologically equivalent to the site occupied by the phosphate group of FMN in flavodoxin. This (proximal) cluster is the closest one to the active site and is located near the enzyme center. The mesial [Fe3S4] and distal [Fe4S4] clusters are bound to the remaining domain that represents 35% of the small subunit and has little secondary structure. This region is missing in some [NiFe]-hydrogenases, such as NAD(P)1-reducing and energy converting enzymes. With the exception of one of the Fe ligands of the distal cluster, that is the Nd atom of a surface exposed histidine side chain, the cluster Fe-atoms are coordinated by cysteine thiolates. The Ni-Fe active site of the D. gigas enzyme is located in the large subunit (Figure 2). Towards the end of the large subunit chain tracing process, we found that there was no electron density corresponding to the last 15 C-terminal residues of the gene sequence [3]. This was surprising because the last visible residue, His536, was deeply buried in the structure. However, shortly thereafter, we noticed a paper by Menon et al. that reported the proteolytic cleavage of a 15-residue C-terminal peptide as part of the maturation process of the large subunit of D. gigas [NiFe]-hydrogenase [12]. In the homologous [NiFeSe]-hydrogenase from Methanococcus voltae, the equivalent of His536 is the C-terminal residue of a 25-residue long third Met. Ions Life Sci. 2009, 6, 151–178

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subunit [13]. Depending on selenium availability, this third subunit can contain either Cys or SeCys as nickel ligand. High resolution structures of the D. vulgaris Miyazaki [4] and D. gigas hydrogenases showed that the C-terminal histidine binds a Mg ion. Upon the cleavage of the C-terminal peptide, the large subunit undergoes a substantial conformational rearrangement burying two C-terminal a-helices. Additional structures of [NiFe]-hydrogenases from D. fructosovorans [5], D. desulfuricans [7], and the [NiFeSe] enzyme from Desulfomicrobium baculatum [6] indicated that all these enzymes have essentially the same domain structure and polypeptide fold. Only the [NiFeSe] enzyme from Dm. baculatum [6], is somewhat different from the Desulfovibrio [NiFe]-hydrogenases: (a) the mesial [Fe3S4] cluster is replaced by a third [Fe4S4] cluster (Figure 2); (b) the Mg ion that binds to the C-terminal histidine of the large subunit in the other enzymes is replaced by an iron ion; (c) near the active site there is a tightly bound H2S molecule, and (d) the Cys530 Ni-ligand of D. gigas is replaced by a seleno-cysteine (SeCys). Additional differences are found at the enzyme surface.

2.2. The Active Site at Medium Resolution In 1993, after having defined the molecular envelope, we calculated a 5 A˚ resolution anomalous scattering difference map of D. gigas hydrogenase. This map showed four peaks more or less linearly arranged and separated by a center-to-center distance of about 12 A˚ [14]. The two [Fe4S4] clusters were assigned to the strongest peaks, whereas the intermediate [Fe3S4] cluster and the active site were considered to generate the third and fourth weaker peaks, respectively. The latter was thought to correspond to a single Ni ion. Higuchi et al. found the same distribution of metal sites in a 4 A˚ resolution analysis of D. vulgaris Miyazaki [NiFe]-hydrogenase [15]. However, when the resolution of the X-ray data from D. gigas enzyme crystals was extended to 2.85 A˚, we realized that the weak anomalous scattering difference peak at the active site corresponded to a different metal ion close to Ni [3]. With the benefit of hindsight we can now conclude that this was to be expected; the anomalous differences we used were obtained using Cu Ka X-ray radiation, the wavelength of which corresponds to the low energy side of the Ni absorption edge. [NiFe]-hydrogenases have two CxxC conserved motifs at the N- and C-terminal ends of the large subunit, respectively. Before the crystal structure was determined, site-directed mutagenesis and EXAFS studies had indicated that the four cysteines of the two conserved motifs coordinated the Ni ion [16]; this was later confirmed by the X-ray analysis [3]. Thus, it was easy to assign the Ni site in the electron density because it was known, from Met. Ions Life Sci. 2009, 6, 151–178


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additional EXAFS analyses of homologous [NiFeSe]-hydrogenases, that it had to be terminally bound to the thiolate of Cys530 [17–19]. The second metal ion was then tentatively assigned to iron because of the reported content of 12 1 iron atoms per hydrogenase heterodimer [3,20]. The active site model at 2.85 A˚ resolution was completed with three ligands, modelled as water molecules, bound to the putative iron ion [3] (X in Figure 3). When in 1996 we started using cryogenic methods and synchrotron radiation with tunable X-ray wavelengths, it became much easier to obtain high resolution, good quality X-ray diffraction data. Crystals could be stored indefinitively in liquid nitrogen and were much more resistant to radiation damage when kept at 100 K during X-ray data collection. Also, complete data sets could be measured much faster from single crystals thanks to the intense synchrotron X-ray beam. Thus, crystallographic evidence for the presence of an active site iron ion was obtained using anomalous scattering X-ray data collected at 1.733 A˚ and 1.750 A˚ (at either side of the iron absorption edge) at the European Synchrotron Radiation Facility

Figure 3. Model of the active site of [NiFe]-hydrogenase from Desulfovibrio gigas at 2.85 A˚ resolution [3]. Three water molecules (Wat) were modeled as ligands of X that, in turn, was refined as an Fe ion. I represents a bridging ligand, it was modeled as an oxygen atom. Met. Ions Life Sci. 2009, 6, 151–178

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(ESRF). A significant peak for the second active site metal ion was only observed in the anomalous difference map calculated from data collected at l ¼ 1.733 A˚ [21]. A similar analysis of D. vulgaris Miyazaki [NiFe]-hydrogenase confirmed this result [4]. In the latter study the assignment of the nickel next to the active site iron was also confirmed crystallographically.

2.3. The Active Site at Higher Resolution A 2.54 A˚ resolution analysis of D. gigas [NiFe]-hydrogenase using a new crystal form and data collected at the ESRF, confirmed the presence of three non-protein ligands to the active site iron ion. However, the shape of the electron density peaks was elongated and could not correspond to monoatomic ligands [21]. Earlier, a FTIR spectroscopic analysis of Chromatium vinosum [NiFe]-hydrogenase (now Allochromatium vinosum) had revealed the existence of three intrinsic high frequency bands that shifted when the redox state of the enzyme was modified [22,23]. When bacteria were grown in a medium containing 13C and 15N, the isotopic effect made these bands shift in a way that indicated they corresponded to two cyanides and one carbon monoxide [24]. The two cyanide bands were found to arise from vibrationally coupled species [25]. Species that were EPR-silent could now also be characterized by FTIR spectroscopy. For instance, an EPR-silent diamagnetic unready oxidized species was identified and designated as Ni-SU. Because equivalent FTIR bands were subsequently obtained with the D. gigas enzyme (Figure 4), we proposed that they originated from the iron ligands, named L1, L2 and L3 that were modeled as diatomic species in our electron density maps (Figure 5) [21]. L3 sat in a hydrophobic pocket and, consequently, it was assigned to CO. On the other hand, L1 and L2 were hydrogen-bonded to the protein (Figure 5), and were modelled as cyanides [26]. Initially, the nature of the diatomic iron ligands was controversial because in the 1.8 A˚ resolution structure of the D. vulgaris Miyazaki hydrogenase a strong electron density peak corresponding to L1 was assigned to SO [4). In order to explain the FTIR spectra, which were essentially identical to those of the other enzymes (S. Albracht, personal communication), L2 and L3 were assigned to a mixture of CO and CN. However, there was no evidence for a SO ligand in the other structures [6,7,21,27], that had very similar electron density for the three diatomic ligands. The strong L1 electron density observed in the first D. vulgaris Miyazaki hydrogenase structure remains unexplained. In fact, recent high-resolution crystal structures of this enzyme have very similar electron densities for L1, L2, and L3 [28,29]. A detailed FTIR study has shown that, indeed, D. vulgaris Miyazaki hydrogenase also contains a Fe(CN)2CO unit [30]. Met. Ions Life Sci. 2009, 6, 151–178


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Figure 4. FTIR spectra of Desulfovibrio gigas [NiFe]-hydrogenase poised at different redox potentials A ¼ – 210 mV , B ¼ – 300 mV, C ¼ – 400 mV, D ¼ – 500 mV, and E ¼ – 600 mV. For details see [26].

There are two coordination sites available for substrate binding at the active site: one that bridges the Ni and Fe ions, called E1 and one that is terminal to the Ni, called E2 (Figure 5). In the initial as-prepared (oxidized, unready) [NiFe]-hydrogenase structures E1 was modeled as occupied by either an oxo ligand [21] or by sulfide [4,7]. These models have been recently corrected, as we and others have found crystallographic evidence showing that in the unready oxidized hydrogenases there is a peroxo, rather than an Met. Ions Life Sci. 2009, 6, 151–178

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Figure 5. The active site of Desulfovibrio gigas [NiFe]-hydrogenase and its environment at 2.56 A˚ resolution [21]. The three diatomic ligands are labeled L1, L2, and L3. Based on their environments L1 was modeled as CO and L2 and L3 as CN (see text).

oxo, ligand bound to E1 [21,31] (see below). On the other hand, the crystals of two reductively activated enzymes did not contain any detectable electron density corresponding to the E1 or E2 sites [6,32]. However, this does not mean that the sites are empty; hydride binding to one or both sites in the reduced active center would not be detected at the resolution of these analyses.

3. HYDROGENASE MATURATION AND ACTIVE SITE ASSEMBLY As stated in the Introduction, hydrogenases are metallo-enzymes that represent one of the most striking examples of convergent evolution; their active site iron ions share the unique feature of being coordinated, in the case Met. Ions Life Sci. 2009, 6, 151–178


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of the [NiFe] and [FeFe] enzymes, with CO and CN and, in [Fe]-hydrogenase, with two CO ligands [2,33]. The synthesis and insertion of these metal centers follow a complex pathway that requires the involvement of accessory proteins with novel biochemical properties.

3.1. Maturation of the Large Subunit The maturation of the large subunit of [NiFe]-hydrogenases requires at least seven proteins. The process can be divided into: (a) synthesis of the apoenzyme, (b) transport and storage of nickel and iron, (c) CO/CN ligand synthesis/binding to iron and partial active site assembly, (d) insertion of nickel, (e) proteolysis of a C-terminal amino acid stretch, and (f) folding of the nascent C-terminal region into the rest of the large subunit. On the other hand, the Fe/S clusters are assembled by the general-purpose Iron sulfur cluster (Isc) scaffold proteins [34]. Hydrogenase synthesis depends on the environment. Under anoxic conditions, Escherichia coli undergoes a major re-organization of its metabolism through the expression of over 300 genes [35]. When hydrogen is present, some microorganisms can detect it thanks to a [NiFe]-hydrogenase-like sensor protein, also called regulatory hydrogenase. Subsequent activation of a kinase and a DNA-binding protein, results in the expression of [NiFe]-hydrogenase structural genes [36]. Both microbial iron and nickel transport are highly regulated in order to avoid Fenton chemistryinduced damage. E. coli and related bacteria have an ATP Binding Cassette (ABC) nickel transporter the corresponding proteins of which are coded by the nik operon. Nickel transport is regulated by NikR, another DNAbinding protein of known three-dimensional structure [37]. Besides relying on specific proteins, iron transport depends on the presence of small molecules called siderophores. We have found that this may also be the case for nickel [38]. In subsequent steps, the active site ligands have to be synthesized and/or ligated to iron and the active site has to be completely assembled and inserted into the large subunit. This process is controlled by the six gene products from the hyp operon and a nickeldependent protease.

3.2. Cyanide, Carbon Monoxide, and Iron Insertion Several years ago, Barrett and coworkers [39] found that inactivation of the enzyme carbamoylphosphate (CP) synthase (PyrA) completely abolished hydrogenase activity in Salmonella typhimurium. However, the basis for this Met. Ions Life Sci. 2009, 6, 151–178

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phenomenon was not further investigated. More recently, Paschos, Glass, and Bo¨ck showed that CN synthesis in E. coli requires two hyp operon gene products, namely, HypE and HypF [40]. HypF is an 82 kDa protein homologous to eukaryotic acylphosphatases at its N-terminal region and to O-carbamoyltransferases at its C-terminal end [41]. The homology of the N-terminal stretch of HypF to acylphosphatases has been confirmed by X-ray crystallography [42]. HypF hydrolyzes CP, transferring the carbamoyl moiety to the C-terminal Cys thiolate of HypE. The formation of a HypF-HypE complex has been shown experimentally [43–45]. HypE is a 35 kDa monomeric protein with amino acid sequence similarities to aminoimidazole ribonucleotide synthase (PurM) and selenophosphate synthase (SelD) [46]. The former catalyzes the dehydration of aminoimidazole ribonucleotide and requires ATP. Its ATP-binding motif is conserved in HypE. The process consists of the following steps: carbamoyl gets transferred by HypF to the HypE C-terminal cysteine, where it is dehydrated generating a HypE-CysSCN complex; next, and in sequential steps, two modified HypEs donate two CN ligands to the nascent Fe unit of hydrogenase active site. It was originally speculated that CP was also the source of CO [40]. However, this notion has been discredited by recent experiments using 13CO2 because only CN was labeled (CO2 is a CP precursor). Since CO was not labeled, its synthesis must follow a different pathway. Roseboom et al. have postulated that CO is synthesized from acetate or one of its precursors [47]. Another possibility is that CO is incorporated directly from the medium without having any specific synthetic pathway. HypC interacts very strongly with both the hydrogenase large subunit and with HypD. HypC is a 9.6 kDa protein with a CxxxP motif in its N-terminal region [48]. HypD is a 41.4 kDa protein that has a [Fe4S4] cluster [47,49]. The HypC-HypD complex is not detected in the presence of citrulline, a CP source. In the absence of nickel, HypC forms a complex with the hydrogenase large subunit HycE. It is thought that the HypC-HypD complex gets the CN ligands from HypE and then transfers them to the hydrogenase, a notion that is comforted by the fact that a HypC-HypD-HypE complex has been characterized [49]. A very recent crystallographic study of HypC, HypD, and HypE has provided a structural interpretation concerning the mechanism by which CN ligands get transfered to the prospective active site Fe (Figure 6). The [Fe4S4] cluster environment in HypD is similar to that of ferredoxin. Although under reducing conditions the complex between HypC and the hydrogenase large subunit precursor preHycE is stable, it dissociates when exposed to alkylating agents. This is consistent with the idea that HypC forms a covalent complex with preHycE by means of its C-terminal cysteine [48]. Nickel also plays a role in the dissociation of the HypC-preHycE complex. Met. Ions Life Sci. 2009, 6, 151–178

Figure 6. Hypothetical cyanation reaction by thiol redox signaling in the maturation of [NiFe]-hydrogenase [93]. The model is based on the crystal structures of the HypC, HypD and HypE.

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3.3. Nickel Insertion Although, as shown by a HypB mutant that lacked hydrogenase activity, HypB is involved in nickel transport/insertion, the enzymatic activity could be restored with added nickel in the growth medium. This suggests that HypB is not essential for nickel insertion [50]. When nickel is replaced by zinc there is no proteolytic maturation showing that the protease is nickelspecific [48,51]. The structure of HypB from M. jannaschii has been reported recently [52]. Nickel insertion in CO dehydrogenase and urease is mediated by similar GTPases [53,54]. Some HypBs may store nickel ions using a histidine-rich stretch at their N termini. In the absence of this region, HypB only binds one Ni ion per protein molecule [55]. Indeed, after the histidinerich sequence, the N-terminal region of E. coli HypB has a high-affinity Ni-binding site with a CxxCxxxxxC sequence [56]. HypA interacts with HypB and it may be the protein that provides nickel to HycE. The HypB GTPase activity may provoke the conformational change needed for nickel insertion by HypA. SlyD is a proline cis/trans isomerase that interacts with HypB. As with the latter, the deletion of the corresponding gene lowers hydrogenase activity but nickel addition abolishes this effect.

3.4. Proteolytic Cleavage of the Large Subunit C-Terminal Extension This process represents the last step in the maturation process [57]. Depending on the hydrogenase, the C-terminal extension can have between five and 32 amino acid residues [58]. Thus, about two-thirds of the C-terminal 25-residue extension of the E. coli hydrogenase 3 can be genetically removed without consequences for either cleavage or subunit maturation. On the oher hand, further truncation strongly reduces precursor stability [58]. The proteolytic enzyme is very specific and there is only one isoform for each hydrogenase in a given organism. The crystal structure of the 17.5 kDa HybD protease is available [59]. Instead of nickel, the structure contains a cadmium ion coordinated by the carboxylate oxygens of glutamic and aspartic acids, the imidazole nitrogen of a histidine and a water molecule. The as-purified HybD does not contain metal [59,60], and in vitro it only cleaves its substrate upon nickel addition. Furthermore, the enzyme activity is insensitive to standard protease inhibitors [61,62]. These observations suggest that HybD recognizes nickel bound to the large subunit precursor and that cadmium replaced physiological nickel in the crystal structure [62]. The maturation endopeptidase Met. Ions Life Sci. 2009, 6, 151–178


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proteolytic site of different hydrogenases can vary being located between either His or Arg and a nonpolar residue such as Met, Ile, Val or Ala. Even this relative conservation is not a strict requirement, as shown by the fact that most mutations at these sites do not prevent proteolysis. It is thus tempting to conclude that the nickel ion, when bound to the large subunit precursor, determines the regiospecificity of the cleavage site [58,62,63].

4. ELECTRON TRANSFER The identity of the hydrogenase physiological redox partner depends on its intracellular location and on its eventual association with another catalytic entity. Examples of such associations are the cytoplasmic bidirectional hydrogen:NAD(P)1 oxidoreductases found in both bacteria and archaea. Monofunctional periplasmic hydrogenases, which are mostly hydrogenoxidizing enzymes, transfer electrons through c-type cytochromes to the cytoplasmic side of the membrane. The resulting protons remain in the periplasm and contribute to the formation of an ATP-generating transmembrane proton gradient. All the available [NiFe(Se)]-hydrogenase structures correspond to periplasmic uptake enzymes of sulfate-reducing bacteria. In enzymes from Desulfovibrio species, electrons move from the active site to the redox partner through a proximal [Fe4S4], a mesial [Fe3S4] and a distal [Fe4S4] cluster (see Figure 2 in Section 2.1). Consecutive redox centers in this pathway are separated by center-to-center distances of about 12 A˚, which are typical for electron transfer [64]. The mesial position of the [Fe3S4] cluster is surprising given the fact that its redox potential is much more positive than that of hydrogen oxidation [65]. Indeed, this cluster should remain reduced during catalysis. It may be that reduction of either the proximal or the distal [Fe4S4] cluster transiently decreases the mesial cluster redox potential making it more compatible with electron transfer in hydrogenase. In order to elucidate the role of the [Fe3S4] mesial cluster we converted it to a [Fe4S4] center in the D. fructosovorans [NiFe]-hydrogenase [66]. The mutated enzyme had wild-type activity indicating that internal electron transfer is not rate-limiting. The reaction rate must then depend on the other processes such as substrate access to the active site, hydrogen heterolytic cleavage, proton transfer from the active site to the solution or electron transfer to a redox partner. Studies with different electron acceptors suggest that the catalytic rate is defined by their interaction with hydrogenase [67]. [NiFe]-hydrogenases are good examples of enzymes having specific redox partner recognition sites. Thus, a crown of acidic residues surrounds the His ligand of the distal [Fe4S4] cluster in different Desulfovibrio species. This Met. Ions Life Sci. 2009, 6, 151–178

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negatively-charged crown is likely to interact with positively-charged patches found near one of the heme groups on both soluble and membranebound c-type cytochromes [3]. The redox complex is probably short-lived, given the binding Km values which are typically micromolar [68]. Electrons are ultimately discarded through the reduction of a varity of cytoplasmic terminal acceptors, such as sulfate in sulfate-reducing bacteria.

5. PROTON TRANSFER Several years ago, [NiFe]-hydrogenase three-dimensional structures were used to propose plausible proton tranfer (PT) pathways. In the 2.8 A˚ resolution structure of an oxidized form of the D. gigas enzyme (pdb entry 1FRV), the side chains of four histidines, two water molecules and a carboxylate group formed a plausible pathway for proton transfer from the Ni-Fe bridging Cys533 ligand to the protein surface [3]. However, this proposition had two problems: a subsequent study at 2.54 A˚ resolution [21] showed that one of these residues, modelled as histidine, was leucine instead (M. Rousset, personal communication). In addition, electron density that bridged the side chains of the C-terminal His residue and a Glu carboxylate was modelled as water but, in reality, it corresponded to the Mg(II) ion discussed above (pdb entry 2FRV). The C-terminal His seems to help stabilizing the C-terminal region, which may be its main role, rather than PT. A very recent study on PT pathways obtained using a quantum mechanical/ molecular mechanical (QM/MM) approach is shown in Figure 7. As mentioned above, in Dm. baculatum [NiFeSe]-hydrogenase the terminal Ni ligand of the D. gigas enzyme, Cys530, is replaced by a SeCys. The 2.1 A˚ resolution crystal structure of the active, reduced form of this enzyme indicated that the Se atom forms a hydrogen bond with the carboxylate of a glutamate side chain, corresponding to Glu18 in D. gigas. This residue is conserved in all [NiFe]-hydrogenases [6]. The SeCys and Glu residues, along with the Ni ion, have temperature (B) factors that are higher than those of atoms in their surroundings. This may be due to a mixture of protonation states of these species in the crystal structure. Similar high B-factors for these two residues have been found in other crystal structures. Consequently, the (Se)Cys and Glu residues are probably constituents of the proton transfer pathway in all [NiFe]-hydrogenases. In fact, in oxidized forms of D. desulfuricans ATCC 27774 [7] and D. fructosovorans [NiFe]-hydrogenases [27] the thiolate of the cysteine residue corresponding to Cys530 in D. gigas displays more than one conformation. There is a precedent for proton transfer from a Ni site to a thiolate ligand in a model compound [69]. Met. Ions Life Sci. 2009, 6, 151–178


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Figure 7. Two possible proton transfer pathways (A and B) in D. fructosovorans [NiFe]-hydrogenase based on a semi-empirical quantum mechanical/molecular mechanical study [94]. Similar results have been recently obtained by Texeira et al. [95].

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The Ni-based EPR spectroscopic properties of the D. fructosovorans enzyme were not modified by the mutation of the glutamate, corresponding to Glu18 in D. gigas, to Gln. However, the mutated enzyme was catalytically inactive and did not perform the H/D isotope exchange. Only the para-H2/ ortho-H2 conversion was still detectable [70]. These results imply that although the mutated enzyme still catalyzed the heterolytic cleavage of H2, the proton/deuterion exchange between solvent and the active site did not take place. Thus, as potulated from the crystallographic results, Glu18 is essential for proton transfer. Matias et al. have proposed a proton transfer pathway based on the crystal structure of oxidized D. desulfuricans ATCC 27774 [NiFe]-hydrogenase [7].

6. OXIDIZED INACTIVE STATES OF THE [NiFe]HYDROGENASE ACTIVE SITE As mentioned above, hydrogenases can be reversibly inactivated by oxygen. Recent revisions and higher resolution oxidized hydrogenase structures have revealed a series of modifications of the thiolates of the cysteine ligands of their active sites. The 1.8 A˚ resolution orthorhombic crystal structure of the as-prepared D. fructosovorans enzyme we reported in 1998 [71] has been recently re-analyzed using quantum refinement and our X-ray diffraction data by So¨derhjelm and Ryde [72]. Their results indicate that Cys68 and Cys530 (D. gigas numbering), were 5% and 20% modified to sulfenic acid, respectively. In addition to this orthorhombic crystal structure, a 1.8 A˚ resolution monoclinic crystal structure of as-prepared hydrogenase was originally published by us in 2002 [27]. More recently, we have revised the structure of this monoclinic form [31] and found that, as in the revised orthorhombic structure described above [72], its active site has both an alternative conformation of Cys530 and the 30% modification of Cys68 to sulfenic acid. Our revision also included the proposition that the bridging E2 site is about 70% occupied by a peroxide species (replacing the oxo ligand we had reported earlier for this site). Similarly, in the recent crystal structure of an unready state of D. vulgaris Miyazaki [NiFe]-hydrogenase, the thiolates of the side chains corresponding to D. gigas Cys68 and Cys530 are 53% and 39% modified to sulfenates, respectively; there is also a 59% occupied NiFe bridging putative peroxo ligand [29]. Thus, the active site seems to be structurally heterogenous in all the crystals containing oxidized unready enzyme. There is a consensus that the oxidized ready Ni-B form has Ni(III) and a bridging hydroxo ligand [31,73,74] (Figure 8A). In our recently revised crystal structure of as-prepared enzyme (Figure 8B), the two atoms of the peroxo ligand do not refine Met. Ions Life Sci. 2009, 6, 151–178


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Figure 8. Different states of the active site of D. fructosovorans [NiFe]-hydrogenase. (A) the putative Ni-B form; (B) putative Ni-A species with a bridging (hydro)peroxo ligand; (C) a sulfenate-containing structure. Results taken from [29] and [31].

to the same occupancy: the oxygen atom closer (proximal) to the Fe ion appears to be more occupied than the distal one. This, in turn, indicates that the proximal site is likely to be occupied by a (hydr)oxo ligand in a significant percentage of hydrogenase molecules. So, when compared to the NiB form, the structures of the unready Ni-A and Ni-SU states appear not to be as well defined. Both the peroxo ligand and cysteine sulfenates (Figure 8C) could in principle be signatures of the unready state. The following reactions may explain the observed structures. At room temperature, the reaction of molecular oxygen with partially reduced hydrogenase is likely to generate a (hydro)peroxo bridging species according to: O2 þ NiðIIÞ þ e þ Hþ ! NiðIIIÞ-OOH

ð2Þ

Taking the redox potentials into account, the electron in reaction (2) is likely to come from the oxidation of the [Fe3S4]0 cluster to [Fe3S4]11. The Met. Ions Life Sci. 2009, 6, 151–178

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resulting (hydro)peroxo ligand can subsequently react with a cysteine thiolate to yield a bridging (hydr)oxo and a sulfenate according to: NiðIIIÞ-OOH þ Cys-S ! NiðIIIÞ-OH þ Cys-SO

ð3Þ

We believe that reaction (3) leads to dead enzyme as suggested by the fact that anaerobically purified hydrogenases are much more active than their reductively re-activated, aerobically purified counterparts [31]. In addition, sulfenic acid formation according to reaction (3) might explain the gradual irreversible enzyme inactivation that is observed in electrochemical studies [F. Armstrong, personal communication]. Indeed, according to the recent DFT analysis of [NiFe]-hydrogenase discussed above, the DG0 of reaction (3) is about –180 kJ/mol [72], which is more than twice as high as the activation energy of + 88 kJ/mol reported for the enzyme in the unready state [26,75]. An -SO ligand will stabilize the lower accessible redox state of Ni, in this case Ni(II), because sulfenate is not as good a s donor as thiolate [76]. We have previously argued that the (hydro)peroxo bridging species is the signature responsible for the unready Ni-A EPR signal [31]. However, the situation is complicated by the fact that (i) ENDOR results from Carepo et al. indicated that after the oxidation of the Ni-C state by air, the labeled oxygen atom of H217O from the solvent medium is bound to Ni in the Ni-A form [77], and (ii) 17O2 also modifies the EPR spectra of the oxidized states of hydrogenases [78]. This leads to the need of an exchange reaction between the putatively O2-generated (hydro)peroxo ligand and water. A possibility is provided by reaction (4) [79] were it is assumed that the exchanging species is Ni-SU, a diamagnetic form, one electron more reduced than Ni-A (Figure 9). Indeed, recent electrochemical results postulate that in the activation process the unready Ni-A state is first reduced to the diamagnetic Ni-SU and subsequently undergoes a slow, reversible step to a transient state [80]. Reaction (4) is analogous to the exchange reaction of peroxide with water in the model heme catalyst, microperoxidase-8 (the Fe(II) ion has been omitted for clarity) [79]: NiðIIÞ-OOH þ e þ Hþ ! NiðIIIÞ-O2 þ H2 O ! NiðIIÞ-OOH þ e þ Hþ

ð4Þ

The (hydro)peroxo-containing species comes from equation (2) and the bridging peroxo ligand will be labeled if this reaction is carried out in the presence of 17O2. Conversely, if activation takes place in the presence of H217O, the labeled water oxygen atom will exchange as in equation (4). The electron would come from the oxidation of [Fe3S4]0. Reaction (4) explains the observed absence of H217O exchange in the Ni-A state [77] Met. Ions Life Sci. 2009, 6, 151–178


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Figure 9. Postulated redox states of the different stable active site intermediates of [NiFe]-hydrogenase. Adapted from [91]. ‘‘?’’ indicates that the bridging site E2 may be occupied in Ni-SI.

because in this form the enzyme has no electrons available to form Ni(III)-O2. In the active site structures of unready hydrogenase, both oxygen atoms of the peroxo ligand are at a binding distance from Ni, which can explain the previous 17O2 EPR and H217O ENDOR spectroscopic results. The possibility of O exchange between H2O and a peroxo ligand at the hydrogenase active site was evoked by Lamle, Albracht, and Armstrong [80]. As a function of their environment, microorganisms have hydrogenases with various degrees of oxygen tolerance. Understanding the structural basis of this resistance is central to ongoing, technologically-oriented, hydrogenase research. To this date, only hydrogenase structures from sulfatereducing bacteria are known. They all have very limited oxygen tolerance. On the other hand, the most remarkable and best characterized oxygenresistant enzymes come from the Ralstonia Knallgas bacteria for which no structure is available. Ralstonia eutropha (now R. cuprivorans) has a membrane-bound [NiFe]-hydrogenase coupled to the respiratory chain and a cytoplasmic soluble enzyme, which oxidizes H2 and reduces NAD1. [81]. The latter is postulated to have two extra CN ligands, one on each of the metal ions of the active site. In addition, the Ni ion would have O/N coordination [82]. The additional CN ligands would be involved in O2 resistance but may dissociate during catalysis. Met. Ions Life Sci. 2009, 6, 151–178

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7. SUBSTRATE BINDING AND CATALYSIS As shown in equation (1) the reaction catalyzed by hydrogenases involves three components: molecular hydrogen, protons, and electrons. In addition, hydrogen cleavage is heterolytic, implying that the active site must also bind hydride. These species are undetectable by X-ray crystallography at the resolutions of the reported studies. Consequently, X-ray crystallography can only provide indirect evidence of substrate binding. Several stable Ni-Fe active site intermediates have been characterized by both EPR and FTIR spectroscopy (see Figures 1 and 4 in Sections 1 and 2.2, respectively). Above, we have addressed the crystallographic data concerning the nature of oxygenic ligands in the active site of unready (Ni-A and Ni-SU) and ready enzyme forms (Ni-B). No exogenous ligands are evident in the crystal structures of reduced, active hydrogenases [6,32]. However, a hydride is likely to occupy the E2 site (see Figure 5 in Section 2.2) making the Ni coordination square pyramidal. Also, the Ni-Fe distance in the reduced enzyme is 2.6 A˚, consistent with the binding of a bridging hydride [83]. Due to electron and proton transfer reactions, different Ni-Fe active site states are at thermodynamic equilibrium making it difficult to obtain hydrogenase crystals in homogeneous states. Theoretical fitting of D. gigas [NiFe]-hydrogenase redox titrations favored the two electron difference model with the active Ni-C form being two electrons more reduced than the oxidized ready Ni-B state [84]. These two EPR active states are considered to be Ni(III) species [85] implying that the two-electron difference arises from the presence of a hydride in Ni-C. Indeed, using HYSCORE and ENDOR spectroscopies, Brecht et al. found direct evidence for bound hydride in the Ni-C state of the regulatory [NiFe]-hydrogenase from R. eutropha [86]. In addition, the g-tensor orientation obtained in a single-crystal EPR study of the Ni-C state of D. vulgaris Miyazaki hydrogenase indicated that the hydride was bound to the Ni-Fe bridging E2 position [87]. This result has been confirmed by a more recent ENDOR/HYSCORE study [88]. Intriguingly, only the Ni-C/Ni-R and one [Fe4S4]21/[Fe4S4] couples were found to be at redox equilibrium with molecular hydrogen [84,89]. This result implies that the enzyme in the hydride-bound Ni-C state can react with hydrogen, and that, consequently, hydride, molecular hydrogens, and protons could occupy both E1 and E2 in a transient form, two electrons more reduced than Ni-C. This transient form would rapidly oxidize to the Ni-R form. There is another observation favoring a putative hydride bound at the bridging E2 site in the Ni-C species and an additional hydrogen binding site at E1: in the crystal structure of D. vulgaris Miyazaki F hydrogenase the added CO inhibitor binds terminally to the Ni at the E1 site [28]. This result agrees with IR spectroscopic data indicating that the vibrational frequency

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of extrinsic bound CO did not correspond to a bridging binding mode in two [NiFe]-hydrogenases [22,90].

8. CONCLUDING REMARKS Hydrogen metabolism is mediated by enzymes that coordinate at least one Fe(CO)x unit. This convergent evolutionary event is remarkable because CO, like CN, is normally toxic to the cells. The diatomic ligands make the iron ion in [NiFe]-hydrogenases both low-spin and redox inactive. It can stay in the 2+ valence state through catalysis thanks to the p-accepting properties of CO and hydrogen-bonded CN. In addition, the CN ligands serve to anchor the iron centers to the protein in both [NiFe]- and [FeFe]hydrogenases. An electron-rich, low spin iron ion has properties that are similar to those of 2nd and 3rd row transition metals, which are known to be good hydrogen catalysts and hydride binders. Thus, Nature has found a cheap solution to hydrogen use and production. Similar approaches may prove useful for the future biotechnological production of this gas as well as its utilization in fuel cells. A more general review on hydrogenases in general can be found in [91].

ACKNOWLEDGMENTS I thank the Commissariat a` l’Energie Atomique and the Centre National de la Recherche Scientifique for institutional support. I also thank Dr. Patricia Amara for critical reading of the manuscript and valuable help with the figures.

ABBREVIATIONS ABC CP DFT ENDOR EPR ESRF EXAFS FMN

ATP-binding cassette carbamoylphosphate density functional theory electron nuclear double resonance electron paramagnetic resonance European Synchrotron Radiation Facility extended X-ray absorption fine structure flavin mononucleotide

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FTIR HYSCORE Isc MIR NAD(P) PT PurM PyrA QM/MM SelD TED

FONTECILLA-CAMPS

Fourier transform infrared hyperfine sublevel correlation iron sulfur cluster multiple isomorphous replacement nicotinamide adenine dinucleotide (phosphate) proton transfer aminoimidazole ribonucleotide synthase carbamoylphosphate synthase quantum mechanical/molecular mechanical selenophosphate synthase two electron difference

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6 Carbon Monoxide and Cyanide Ligands in the Active Site of [FeFe]-Hydrogenases John W. Peters Montana State University, Department of Chemistry and Biochemistry and the Astrobiology Biogeocatalysis Research Center, Bozeman, MT 59717, USA <[email protected]>

ABSTRACT 1. INTRODUCTION 2. [FeFe]-HYDROGENASE STRUCTURE 2.1. [FeFe]-Hydrogenase Overall Structure 2.2. [FeFe]-Hydrogenase Active Site H-Cluster Structure 2.2.1. Biologically Unique Ligands 2.2.2. Ligand Exchangeable Site 2.2.3. Carbon Monoxide Inhibition 2.2.4. Non-Protein Dithiolate Composition 3. [FeFe]-HYDROGENASE SPECTROSCOPIC STUDIES 3.1. Electron Paramagnetic Resonance Spectroscopy and Mo¨ssbauer Spectroscopy 3.2. Infrared Spectroscopy 4. H-CLUSTER MODEL COMPLEXES 4.1. 2Fe Subsite Model Synthesis 4.2. Advanced 2Fe Subsite and H-Cluster Model Synthesis 5. H-CLUSTER BIOSYNTHESIS 5.1. Relevance to Prebiotic Chemistry 5.2. Identification of Genes 5.3. Gene Clusters and Operons


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5.4. Radical-S-Adenosylmethionine Chemistry and Potential Biological/Biochemical Sources 6. FUTURE DIRECTIONS ACKNOWLEDGMENTS ABBREVIATIONS AND DEFINITIONS REFERENCES

PETERS

202 206 207 208 208

ABSTRACT: The [FeFe]-hydrogenases, although share common features when compared to other metal containing hydrogenases, clearly have independent evolutionary origins. Examples of [FeFe]-hydrogenases have been characterized in detail by biochemical and spectroscopic approaches and the high resolution structures of two examples have been determined. The active site H-cluster is a complex bridged metal assembly in which a [4Fe-4S] cubane is bridged to a 2Fe subcluster with unique non-protein ligands including carbon monoxide, cyanide, and a five carbon dithiolate. Carbon monoxide and cyanide ligands as a component of a native active metal center is a property unique to the metal containing hydrogenases and there has been considerable attention to the characterization of the H-cluster at the level of electronic structure and mechanism as well as to defining the biological means to synthesize such a unique metal cluster. The chapter describes the structural architecture of [FeFe]-hydrogenases and key spectroscopic observations that have afforded the field with a fundamental basis for understanding the relationship between structure and reactivity of the H-cluster. In addition, the results and ideas concerning the topic of H-cluster biosynthesis as an emerging and fascinating area of research, effectively reinforcing the potential linkage between ironsulfur biochemistry to the role of iron-sulfur minerals in prebiotic chemistry and the origin of life. KEYWORDS: hydrogenase hydrogen evolution hydrogen oxidation iron-sulfur enzymes iron-sulfur cluster biosynthesis nitrogenase nitrogen fixation non-protein ligands prebiotic chemistry proton reduction

1. INTRODUCTION The presence of both carbon monoxide and cyanide as ligands to Fe atoms at the active site of the [NiFe]- and [FeFe]-hydrogenases is still a surprise to many even after more than ten years since the first definite evidence was presented on their existence [1]. To date, no enzymes other than the aforementioned hydrogenases have been shown to possess both ligands to Fe in the active form. The three classes of hydrogenases, described in this chapter [FeFe] (previously termed Fe-only hydrogenases), the preceding chapter [NiFe], and the next chapter [Fe], share in common carbon monoxide ligation to Fe atoms and also have common structural features and reactivity [2–28]. Despite this, however, it seems clear that these three classes of enzymes have separate evolutionary origins, and common structural and functional features have arisen by divergent evolution [29–33]. These Met. Ions Life Sci. 2009, 6, 179–218

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inferences can be attributed to the lack of primary sequence similarity between the three classes and distinct differences in the occurrence of these enzymes in various microorganisms. The aforementioned hydrogenases occur only in microorganisms where they function in microbial metabolism to either couple hydrogen oxidation to energy-yielding processes or to dispose of excess electrons that accumulate during anaerobic metabolism [2,3]. Amongst microorganisms, these hydrogenases have been found to occur in bacteria, archaea, and lower eukaryotes including protists and algae. Analysis of available genomes indicates that the [NiFe]-hydrogenases are widely spread in bacteria and archaea and are prevalent in cyanobacteria but have yet to be observed to occur in any eukaryotes [29–31,33]. In contrast, [FeFe]-hydrogenases are widely distributed among anaerobic and facultative anaerobic bacteria, some protists and algae, but do not occur in the genomes of cyanobacteria or archaea [30–33]. A third class of hydrogenase exclusive to methanogens, termed [Fe]-hydrogenases, occurs with an iron carbonyl in association with an organic cofactor [5,28,34] (Chapter 7). Regardless of evolutionary origin, it is rational to think that one or more of the [Fe]-, [FeFe]-, and/or [NiFe]hydrogenases have an ancient earth origin given the potential roles for iron-sulfur minerals and derivatives in prebiotic chemistry [35–43] and the presumed importance of reversible hydrogen oxidation chemistry in early energetics [44]. Further examination of the phylogeny of organisms in which these enzymes occur will provide insights into the origins of these enzymes. The unique structural character of the carbon monoxide in all three classes [1,12,14,21,45–59] and cyanide in the [FeFe]- [12,21,48,51,52,57] and [NiFe]-hydrogenases [1,12,14,,45–51,53,54,56,58,59] distinguishes these enzyme active sites from other organometallic cofactors found in nature [60] and makes these enzymes of significant interest to basic scientists from evolutionary biologists to physical chemists. In addition, the relevance to prebiotic chemistry [60–63] coupled with the potential for the enzymes as Noble metal-free solutions to alternative and renewable energy [64–74] may suggest that the studies on the structure, function, biosynthesis, and origin of hydrogenases may hold both the keys to the origin of life as well as the future of life on earth.

2. [FeFe]-HYDROGENASE STRUCTURE 2.1. [FeFe]-Hydrogenase Overall Structure In the late 1990s, the structure of [FeFe]-hydrogenases from two different microbial sources (Clostridium pasteurianum and Desulfovibrio desulfuricans) Met. Ions Life Sci. 2009, 6, 179–218

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were revealed [11,18]. The [FeFe]-hydrogenase from Clostridium pasteurianum (CpI) determined by our group is a B60 kD monomeric enzyme that is localized in the cytoplasm and functions in the coupling of proton reduction to the oxidation of reduced electron carriers that accumulate during carbohydrate fermentation [11]. The Desulfovibrio desulfuricans [FeFe]-hydrogenase (DdH) determined by the Fontecilla-Camps group is a dimeric enzyme, with a large and a small subunit, is localized in the periplasm and consequentially is presumably involved physiologically in hydrogen oxidation [18]. In addition to the differences in the quaternary structure and cellular localization of the C. pasteurianum and D. desufuricans [FeFe]-hydrogenases, the enzymes also differ in overall size and in their respective complement of accessory FeS clusters. The C. pasteurianum [FeFe]-hydrogenase exists with an active site H-cluster and four additional FeS clusters including three [4Fe-4S] clusters and a [2Fe-2S] cluster (Figure 1A) while the D. desufuricans enzyme exists with an accessory cluster composition of just two [4Fe-4S] clusters in addition to the active site H-cluster (Figure 1C). In general, all [FeFe]-hydrogenases that have either been characterized biochemically [2,75–91] or simply implicated by searches of deduced amino acid sequences in the genomes of various organisms [30,32,33] share a common architecture with most apparent differences on the complement of various FeS clusters that link the active site H-cluster for reversible hydrogen oxidation to various external electron donors and acceptors. These differences presumably affect the differences in the physiology of organisms that harbor these enzymes by accommodating specific and distinct external electron donors and/or acceptors. The C. pasteurianum monomeric [FeFe]-hydrogenase exists with an overall mushroom shape with a large domain represented by the mushroom cap in which the active site H-cluster resides (Figure 1A and 1B). In addition to the larger H-cluster domain, three other smaller domains can be assigned according to the structural similarity to known ferredoxin proteins [92,93]. The domain proximal to the H-cluster domain contains two [4Fe-4S] clusters and is similar in overall structure to the numerous two [4Fe-4S] cluster-containing ferredoxins for which the structures of several are known (Figure 1A, green domain) [93–98]. This domain is common to many [FeFe]-hydrogenases [18,99] and is also found as a domain in additional redox proteins and enzymes involved for example in respiration and photosynthesis [100,101]. The ferredoxins and the structurally analogous domains are easily distinguished in searches of deduced amino acid sequences in genomes because they possess a well-defined and conserved arrangement of cysteine ligands (eight conserved cysteine residues) that function as covalent thiolate ligands to the [4Fe-4S] clusters. The far N-terminal domain contains a [2Fe-2S] Met. Ions Life Sci. 2009, 6, 179–218


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Figure 1. A. Cartoon depiction of the X-ray crystal structure of CpI [11] showing the overall fold and division into 4 separate structural domains based on FeS cluster content and structural similarity to known ferredoxin proteins. The active site Hcluster domain is shown in blue, the 2 [4Fe-4S] cluster ferredoxin-like domain is shown in green, the N-terminal [2Fe-2S] cluster ferredoxin-like domain is shown in light magenta, the single [4Fe-4S] domain is shown in purple, and the C-terminal region is shown in red. The FeS clusters and CO and CN ligands are shown as space filling models (Fe: dark red, S: orange, O: red, N: blue). B. Zoom view of the active site domain showing the H-cluster cleft formed by the pseudosymmetric symmetry of the two twisted b-sheets on either side. The C-terminus is perpendicular to the bsheets and wraps around the cleft as occupied by the H-cluster. C. Cartoon depiction of the X-ray crystal structure of DdH [18] where equivalent structural regions to CpI are shown in the same colors. D. Cartoon depiction of the homology structure of C. reinhardtii to CpI as determined by the homology server Phyre [225,226]. The green algae C. reinhardtii lacks additional accessory cluster domains and is represented by the homologous active site domain only.

cluster and is also structurally analogous to a class of what is termed planttype ferredoxins for which structures have been described (Figure 1A, magenta domain) [92,102–109]. Again, a characteristic motif of cysteine residues facilitates the identification of structurally analogous domains or ferredoxins of this type in deduced amino acid sequences. An additional small domain containing a single [4Fe-4S] cluster separates the two ferredoxin-like domains (Figure 1A, purple domain). This [4Fe-4S] cluster possesses a unique set of coordinating ligands with the combination of three cysteine thiolates and secondary amine of a histidine imidazole side chain. This combination of ligands in an accessory cluster is a feature only observed in this [FeFe]-hydrogenase and in the characterized structures of Met. Ions Life Sci. 2009, 6, 179–218

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the [NiFe]-hydrogenases from sulfate-reducing bacteria [4,7,8,16,110] but to my knowledge not observed elsewhere in biology. The specific biochemical and physiological role of this unique cluster is not known, but it is possible that the ligand arrangement would contribute to defining the oxidation-reduction potential of the cluster and the presence of the histidine coordination may contribute to proton-coupled electron transfer. In comparison, the D. desulfuricans structure lacks the histidine-coordinating cluster, as well as the [2Fe-2S] clusters and their associated domains (Figure 1C) [18]. The complement of only the additional two [4Fe-4S] cluster containing ferredoxin-like domains (Figure 1, green domains) is observed in many [FeFe]-hydrogenases [18,99] and phylogenetic analysis [30,32,33] reveals this feature in the most deeply rooted organisms suggesting that this arrangement of domains may be ancestral. The aforementioned [FeFe]-hydrogenase architectures (those with either four or two accessory clusters) represent the majority of known [FeFe]hydrogenases. However, as mentioned above, the specific complement of accessory clusters and their associated domains is a distinguishing property and a notable additional variation on this theme observed in [FeFe]-hydrogenases of thermophiles [111], hyperthermophiles [112], and eukaryotes including a group that include the [FeFe]-hydrogenases observed in algae [113–119], which lack accessory clusters entirely (Figure 1D) and other more complex versions occur in the hydrogenosomes of the ciliate Nyctotherus ovalis in which the [FeFe]-hydrogenase functionality is fused to domains that have primary sequence similarities to electron transport chain components [120]. This arrangement implies that the latter [FeFe]hydrogenase can directly reoxidize NADH and has been suggested to support the idea that the hydrogenosome evolved from ciliate mitochondrion [121]. Given the relative position of the accessory clusters of the C. pasteurianum [FeFe]-hydrogenase, a single pathway for the transfer of electrons to and from the active site H-cluster and external donors and acceptors cannot be rationalized [11]. The histidine-ligated [4Fe-4S] cluster and the [2Fe-2S] cluster are located close to the protein surface at the termini of what could be a branched pathway providing the means to interact with electron transfer partners with different chemical characters and may serve as some sort of metabolic switch for the enzymes between hydrogen production and hydrogen oxidation. An assessment of the relative distribution of charge on the surface of the C. pasteurianum [FeFe]-hydrogenase indicates that the protein surface surrounding is distinctly positively charged and in contrast the region surrounding the [2Fe-2S] cluster is distinctly negatively charged which may support interactions with multiple external electron transfer partners. Alternatively, the histidine-coordinated cluster may have a different role that has yet to be determined. Met. Ions Life Sci. 2009, 6, 179–218


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The active site domain is the largest domain in [FeFe]-hydrogenases (B300 amino acid residues) and exists as a pseudosymmetric structure in which two twisted b-sheet structures come together to form a cleft in which the H-cluster is located (Figure 1A and 1B). Covalent coordination of the 6Fe H-cluster to this domain is provided by four cysteine thiolates, two each provided from each pseudosymmetric half. The residues involved in covalent coordination and the associated residues within the environment of the H-cluster are highly conserved [30] providing the basis for a primary sequence signature for the identification of [FeFe]-hydrogenases from genome sequences (Figure 2). These motifs also provide the basis of differentiating between bona fide [FeFe]-hydrogenases and related sequences including eukaryotic narF gene sequences [122–124]. For the C. pasteurianum, the cleft where the H-cluster is located is covered by the C-terminal region wrapping around the cleft perpendicular to the approximate axis of the pseudosymmetric structure (Figure 1A and 1B). In the dimeric D. desulfuricans structure, the region analogous to the C-terminus of C. pasteurianum [FeFe]-hydrogenase is provided by the small subunit (Figure 1C). The small subunit (b-subunit) also encodes the determinants that direct the enzyme to the twin arginine transporter for localization of the enzyme in the periplasm.

2.2. [FeFe]-Hydrogenase Active Site H-Cluster Structure Much of the attention in the structures of [FeFe]-hydrogenases can be correlated to the interest in the structure of the active site H-cluster. The surprises realized in the early structural characterization of the [NiFe]-hydrogenase active sites [4,7,8,16] coupled with the inability to come to a structural consensus based on spectroscopic characterization placed an imperative on defining the structural details of the H-cluster in the late 1990s. In 1996, in a seminal review on iron-sulfur enzymes [60], authored by Richard Holm of Harvard and Edward Solomon together with his postdoctoral associate at the time, Pierre Kennepohl, of Stanford University, highlighted this imperative and described the H-cluster as the most conspicuously undescribed complex iron-sulfur cluster of the time. With the presence of the carbon monoxide and cyanide ligands observed in the [NiFe]-hydrogenase active site [1,12,14,45–50], it was envisioned that this complement of ligands could also be present in the [FeFe]-hydrogenase, but this had not been definitely determined until after the first structures were revealed. The CpI structure, which was described first [11], was originally refined to 1.8 A˚ resolution and afforded sufficient information to make tentative assignments to most of the features of the cluster. The H-cluster in the CpI Met. Ions Life Sci. 2009, 6, 179–218

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Figure 2. Sequence alignments of D. desulfuricans (DdH), T. maritima (Tma), and C. reinhardtii (Crl) relative to C. pasteurianum (CpI). Residues in black are conserved or very similar and residues in gray are not conserved. The secondary structure of CpI is represented by the cartoon below its sequence: red waves (a-helices), blue arrows (b-sheets), yellow semi-circle (turns) and black line (loops). Met. Ions Life Sci. 2009, 6, 179–218


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Figure 3. Schematic of the [FeFe]-hydrogenase active site (A) from C. pasteurianum (CpI) [11] and (B) [NiFe]-hydrogenase active site from D. gigas in the oxidized state [4,47]. For CpI, the unknown atom of the dithiolate ligand is labeled ‘‘X’’ and shown in magenta.

structure (Figure 3A) was described as a [4Fe-4S] subcluster coordinated to the enzyme via four cysteine ligands, one of which serves as a bridging ligand to a 2Fe subcluster in which the diatomic ligands carbon monoxide and cyanide were assigned based on the analogy to the [NiFe]-hydrogenase where these ligands were demonstrated using FTIR spectroscopy (Figure 3B) [1,45,46]. Although FTIR analysis of the [FeFe]-hydrogenase was not available when the structures were determined, it was available soon after providing additional details into the nature of these ligands (discussed in detail below).

2.2.1. Biologically Unique Ligands One of the most surprising features of the structure of the 2Fe subcluster of the H-cluster was the absence of any protein coordination linking the Fe atoms. Unlike the bimetallic active site cluster of the [NiFe]-hydrogenase which exists with a Ni ion linked via cysteine thiolates to a carbon monoxide and cyanide coordinated Fe ion (Figure 3B), the 2Fe subcluster of the H-cluster consists of carbon monoxide and cyanide coordinated Fe atoms bridged by a diatomic ligand assigned as carbon monoxide and a dithiolate ligand of unknown composition (Figure 3A). In the original CpI structure we simply described the dithiolate ligand as a covalent linkage of light atoms since there was no experimental data available for assignment of the composition of this ligand. Very soon after the structure of the CpI enzyme was reported, the DdH structure [18] was determined which afforded a slightly different structure of the H-cluster. As in the case of the CpI structure, the DdH structure was refined to reasonably high resolution allowing ligand assignments at a high level of detail and certainty. It is presumed that both enzymes possess the same prosthetic groups structure and thus the observed differences were attributed to alternative interpretations of some structural features and Met. Ions Life Sci. 2009, 6, 179–218

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differences in states in which the enzyme was poised or trapped during structure determination. With respect to the former, key differences in interpretation were offered at the site of the non-protein bridging ligands of the 2Fe subcluster. In place of a bridging carbonyl, an asymmetric bridging water or hydroxide ligand was assigned. This difference in interpretation was later resolved to be a bridging carbonyl by FTIR spectroscopy and inability to assign a carbonyl ligand at this position in the crystallographic work was attributed to potentially mixtures of oxidation states of the cluster present in the crystals [21]. With regard to the composition of the dithiolate ligand, the Fontecilla-Camps group assigned the ligand as a propane dithiolate (PDT). This is consistent with covalent linkage of light atoms in the CpI structure, however, no experimental evidence was offered and the assignment was largely based on the common use of this ligand in chemical synthesis. The evolution of thoughts on the composition of the dithiolate ligand driven largely by mechanistic considerations is ongoing and quite interesting and will be discussed in detail below.

2.2.2. Ligand Exchangeable Site A third difference observed in the CpI and DdH H-cluster structures is the ligand composition and the coordination environment of distal Fe atom to the [4Fe-4S] cluster. In the CpI structure, the Fe atom is in an octahedral coordination environment with and in addition to the previously mentioned five ligands (the carbon monoxides and cyanide and the sulfur atoms of the dithiolate ligand) significant electron density existed and a weakly bound water molecule was assigned. In the DdH structure, electron density was not observed at the position where the water molecule was assigned in CpI such that the DdH structure was modeled with a vacant coordination site and a Fe atom in a square pyramidal geometry. Shortly after the two structures were described, our group teamed with the Fontecilla-Camps group in an effort to explain these differences. In a review article published in TIBS in 1999 [20], the differences were explained to potentially arise from real differences in oxidation state and evaluated together thought to be mechanistically insightful. In this work, we evaluated the differences in which the enzyme was crystallized prior to the structure determination. For CpI, crystals were grown in a Vacuum Atmospheres Anaerobic Chamber (Cu-containing oxygen consuming catalyst) over a two to three week period in a buffer that contained sodium dithionite at a 1 mM concentration at the start of crystal growth. For DdH the crystals were grown in a Coy Anaerobic Chamber which in contrast to the Vacuum Atmospheres Anaerobic Chamber uses a palladium-based system which catalyzes the conversion of hydrogen and oxygen to yield water. Met. Ions Life Sci. 2009, 6, 179–218


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Figure 4. Summary of the spectroscopic data collected for the five observed physical air , Htrans, Hox, Hred, and HoxCO). For the Hox [11], Hred states of the H-cluster (Hox [18], and HoxCO [19] states the X-ray crystal structures have been determined and are represented as ball and stick models (coloring scheme: Fe, rust brown; S, orange; O, red; N, blue; C, dark gray; unknown atom of dithiolate ligand, magenta). The reported spectroscopic data for Hox and HoxCO is for the CpI enzyme, the Hred state air for DdH, and the Hox and Htrans data is for DvH, which is presumed to be identical to DdH. The oxidation states for the [4Fe-4S] cluster and 2Fe subcluster represent Mo¨ssbauer data, DFT data, and the generally accepted conclusion that the clusters exist in low oxidation states. For the IR data, the band correlated to the bridging CO ligand is in bold and is observed to shift from bridging to partial terminal upon reduction of the Hox state to Hred state. The CX band for the Hred state is intermediate to both CO and CN ligands and cannot be assigned unambiguously to either carbon monoxide or cyanide.

For CpI under the conditions described above the hydrogenase would have most likely consumed the sodium dithionite added to the crystallization conditions over the course of the two to three week incubation period at room temperature required for crystal growth. It is logical to assume that the CpI observed in the crystal structure is poised in the oxidized state (see Figure 4). In contrast, since DdH crystals were grown in the Coy Anaerobic Chamber in the presence of a hydrogen partial pressure, it is reasonable to assume that the enzyme observed in this structure is likely to represent the reduced state of the hydrogenase or a state in which a hydride or hydrogen is bound at the active site (see Figure 4). Taking these considerations into Met. Ions Life Sci. 2009, 6, 179–218

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account, a mechanistic scheme can be proposed in which this ligandexchangeable site is occupied by a water molecule in the oxidized state but when the metal cluster is reduced or hydrogen is available as a coordinating group, water is displaced (see Figure 4). In the DdH structure, it is unlikely that this site is a vacant site but that it is occupied by either hydrogen or a terminal hydride which cannot be resolved by X-ray crystallography.

2.2.3. Carbon Monoxide Inhibition Several subsequent studies involving X-ray diffraction studies and FTIR strongly support the presence of this ligand-exchangeable site at the distal Fe atom of the 2Fe subcluster relatative to the [4Fe-4S] cluster [19,21,52,125]. Interestingly, as in the case of most [NiFe]-hydrogenases [23,50,126] the [FeFe]-hydrogenases are sensitive to inhibition by carbon monoxide [2,91,127–130]. The application of FTIR spectroscopy to the analysis of hydrogenases was really prompted by the observation that the enzymes were inhibited by carbon monoxide and if it was not for this connection we may still be searching for the identity of the diatomic non-protein ligands in these enzymes. We were able to obtain crystals of CpI grown in the presence of carbon monoxide in the same crystal form as native CpI allowing for the analysis of carbon monoxide binding and inhibition by crystallographic difference Fourier analysis [19]. The approach affords a relatively unbiased method to compare different enzyme forms and involves subtracting two X-ray diffraction data sets from one another. Applied to CpI, the approach revealed unambiguously that carbon monoxide inhibits [FeFe]-hydrogenases by binding to the aforementioned ligand-exchangeable site. These results strongly supported the mechanistic inferences derived from the previous structures of the native structures of CpI and DdH. The results could not be immediately reconciled with previous studies and subsequently an additional structure of DdH from the Fontecilla-Camps group was presented as well [21]. This structure was reported to represent a more highly populated structure of the reduced state. In this work, the bridging carbon monoxide ligand was observed in an asymmetric configuration more closely coordinated to the Fe atom of the 2Fe subcluster with the ligand exchangeable site. The modulation in the distance is reasonable given the trans effect of the bridging carbon monoxide coordinated to a position trans of the ligand exchangeable site of the distal Fe. This trans effect is likely critical to the maintenance of the ligand-exchangeable site and the efficacy of the H-cluster as a catalysts in reversible hydrogen oxidation. This is highlighted by our work monitoring the selective photochemical cleavage of inhibitory carbon monoxide by X-ray crystallography [19]. In this experiment, data was collected on the carbon monoxide-inhibited form Met. Ions Life Sci. 2009, 6, 179–218


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of CpI before and during continuous exposure of the crystal to a He/Ne laser source. The two data sets generated (before and during illumination) were examined by difference Fourier methods and since the two data sets differed only by light exposure, subtle differences were distinguishable. The experiment showed that the inhibitory carbon monoxide was selectively cleaved by the presence of a peak of negative electron density at the terminal site when the illuminated X-ray diffraction data was subtracted from the data collected prior to illumination. The data did not show a corresponding positive peak of electron density probably indicating that upon photolytic cleavage, carbon monoxide does not migrate to a specific position. Additionally, electron density features in this analysis provided for a structural affirmation of the importance of the trans effect in maintaining a ligand exchangeable site in the [FeFe]-hydrogenases. Additional electron density features (both positive and negative) surrounding the Fe atom indicate that upon photochemical cleavage the trans effect is relieved allowing the bond distance between the Fe and the bridging carbon monoxide to become shorter. The results suggest that presence of the strong back-bonding ligand trans from the site of reversible hydrogen oxidation makes the site more labile and enhances the ability of this site to cycle dynamically.

2.2.4. Non-Protein Dithiolate Composition As mentioned above, one of the most interesting remaining questions with regard to the structure of the H-cluster and mechanism of reversible hydrogen oxidation at this site is the composition of the non-protein dithiolate ligand of the H-cluster. Although PDT was assigned in the DdH H-cluster [18], the proposal was revised shortly after by the FontecillaCamps group when it was envisioned that the presence of a secondary amine group at the central position of the five atom ligand could potentially act as a catalytic base cycling between protonation states [21]. The presence of a proton donor acceptor group in close proximity to the specific coordination site for reversible hydrogen oxidation was likened to the role proposed for the Ni ion in [NiFe]-hydrogenases. The mechanistic implications of the amine group are potentially very exciting with regard to heterolytic cleavage of dihydrogen [131–133] and provided new targets for synthetic chemists (discussed later), however, there has yet to be any direct chemical or biochemical evidence to support this assignment. Recently, our group has begun to analyze the structure of CpI at B1.4 A˚ resolution [134]. The H-cluster in this structure is clearly resolved, however, refinement even at this resolution does not afford the basis for discriminating the composition of isoelectronic groups. It is generally accepted that the groups adjacent to the thiolate sulfurs by definition are methylenes Met. Ions Life Sci. 2009, 6, 179–218

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and really the group in question is the central group. The group could presumably be an additional methylene, a secondary amine group, or an ether. In an effort to extend the resolution of our recent X-ray crystallographic study, we have used density functional theory to evaluate which of the three aforementioned scenarios is closer to an energy minimum when placed in our refined crystallographic coordinates [134]. A relatively simple approach complicated only by the extent of the model used for the calculations which encompassed a greater than 3.0 A˚ environment of the entire H-cluster was taken. For these studies a methylene (PDT), protonated and nonprotonated amine (DTN), in addition to ether and thioether groups (DTO) were treated as separate optimization runs in the analysis and it was found that an ether group reached optimization with a smaller energy difference. In addition, the structure of the protonated amine group was unstable during the optimization process such that the amine group rests in a position in which a hydrogen bond forms between the amine group and the thiolate group of the subcluster bridging cysteine. This analysis is clearly not unequivocal but it does indicate that the community should keep an open mind with regard to the composition of the ligand and places an imperative on obtaining direct biochemical evidence for the composition of this ligand.

3. [FeFe]-HYDROGENASE SPECTROSCOPIC STUDIES 3.1. Electron Paramagnetic Resonance Spectroscopy and Mo¨ssbauer Spectroscopy Key information regarding the physical features of the H-cluster, spin density, oxidation states of the [4Fe-4S] cluster and 2Fe subcluster, and CO inhibition have been analyzed by EPR [83,88–90,119,125,129,130,135–138], ENDOR [139–141], ESEEM [92,140,142,143], IR [12,21,48,51,52,57], and Mo¨ssbauer spectroscopies [15,144,145]. Of the two catalytically relevant redox states of the H-cluster, Hox is paragmagnetic and Hred is EPR-silent. The Hox state displays a characteristic rhombic EPR signal (S ¼ 1/2, g ¼ 2.10, 2.04, 2.00) attributed to the H-cluster (Figure 4) [88,125]. As discussed in the description of the crystallographic work, the [FeFe]-hydrogenases are sensitive to CO inhibition [2,91,127–130] and for CpI it has been observed that CO inhibition occurs by the binding of CO at the ligand-exchangeable site of the distal Fe of the 2Fe subcluster [19]. CO inhibition of the Hox state, for CpI, DdH, and D. vulgaris Hildenborough (DvH), results in the disappearance of the characteristic rhombic signal of the Hox state and the appearance of a distinct axial EPR signal (g> ¼ 2.01, Met. Ions Life Sci. 2009, 6, 179–218


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g|| ¼ 2.07) characteristic of the HoxCO state [90,92,125]. The distinctly different character of the Hox and HoxCO EPR signal reflects the large difference in cluster symmetry in the two states. EPR is an effective tool for monitoring CO inhibition, for example the aforementioned crystallography experiments involving selective photolytic cleavage of the inhibitory carbon monoxide upon hv irradiation can be monitored directly by the reappearance of the Hox rhombic signal [12,57,89,138]. Unlike CpI [11], DdH, and DvH hydrogenases can be purified aerobically [86,91,146] and sequentially activated under reducing conditions. Because DdH and DvH have identical primary sequences, it is presumed that DvH has the same tertiary structure as DdH [91,99,147] although its structure determination by X-ray crystallography has yet to be reported. The aerobically isolated DdH and DvH are distinct from the aforementioned air , which is presumed catalytically relevant oxidation states and termed Hox air catalytically inactive and EPR-silent [88]. One electron reduction of the Hox species results in an EPR-active (S ¼ 1/2) transitional state (Htrans) [88]. Mo¨ssbauer studies have shown that the Htrans results from the single air electron reduction of the [4Fe-4S]21 subcluster of the H-cluster of Hox [145]. This is supported by the EPR data, which show the appearance of a rhombic EPR signal with g-values similar to those for a typical [4Fe-4S]11 air cluster upon reduction of Hox [88]. The [4Fe-4S] cluster remains in the 21 form for all other H-cluster states [15,145]. EPR-silent [4Fe-4S] Mo¨ssbauer [15,144,145] and additional DFT studies [133,148–150] of the Hcluster and synthetic models [151–155] of the 2Fe subcluster have been directed at trying to gain insights into the oxidation states of the Fe atoms of the 2Fe subcluster in the aforementioned characterized states of the H-cluster. air Þ, Initial studies suggested that the oxidation states were FeIIIFeIII ðHox III III II III II III II II (Htrans), Fe Fe (Hox), Fe Fe (HoxCO), Fe Fe (Hred) Fe Fe air Þ, FeIIFeII (Htrans), [15,144,145]. However, it was noted that FeIIFeII ðHox I II I II I I Fe Fe (Hox), Fe Fe (HoxCO), Fe Fe (Hred) would also fit the data [15,144,145]. More detailed computational studies addressing the issue of H-cluster oxidation states (discussed in detail in Chapter 12) have led to the general accepted view that the 2Fe subcluster exists in the latter, lower oxidation states consistent with the presence of these unique electron donating carbon monoxide/cyanide ligands [133,148–150].

3.2. Infrared Spectroscopy The potential for the presence of non-protein ligands in the H-cluster was realized in the early 90s when nitrogen modulations were observed in ESEEM studies on CpI that were distinct from those that would be Met. Ions Life Sci. 2009, 6, 179–218

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anticipated from protein components [140,143]. As described above, the assignment of carbon monoxide and cyanide as ligands in hydrogenase did not occur until the late 90s from the results of an IR analysis of these enzymes [12,21,48,51,52,57]. The Hox state of CpI can be characterized by five FTIR indicated bands at appropriate stretching frequencies that can be attributed to the carbon monoxide and cyanide ligands [52]. The five bands have been assigned to two terminal carbon monoxide ligands (1948 and 1971 cm1), two terminal cyanide ligands (2072 and 2086 cm1) and a single bridging carbon monoxide (1802 cm1). The Hox state is presumed to correspond as the CpI structure in which a water molecule is coordinated to the ligand-exchangable [4Fe-4S] cluster distal Fe atom of the 2Fe subcluster [156]. FTIR studies of the Hox state of DdH and DvH have also been reported and reveal similar bands [12,21,48,51]. The reduced state of CpI has not been studied in detailed, however, for DdH and DvH, IR characterization has provided some insights into redox-dependent structural changes that occur within the H-cluster during catalysis. These studies show that the IR band attributed to the bridging carbon monoxide is significantly shifted to lower energy from 1802 cm1 to 1894 cm1 upon reduction indicating a change in the character of this bridging ligand to a more asymmetric ligand [12,21,51]. This is supported by the crystallographic characterization of the Hred of DdH as described above having a very asymmetric bridging carbon monoxide ligand that approaches terminal coordination to the [4Fe-4S] cluster distal Fe atom of the 2Fe subcluster [21]. The only other state that has been characterized both crystallographically and by FTIR is the carbon monoxide inhibited state of CpI [19,52]. In this state the bridging carbon monoxide is observed to be more or less symmetric in the crystal structure [19] and in the FTIR the band is observed at 1810 cm1 which is very similar to that observed in the oxidized air and Htrans states have not been characterized state (1802 cm1) [52]. The Hox by X-ray crystallography but from the perspective of the IR of the bridging carbon monoxide, the states display IR bands of 1848 cm1 and 1836 cm1 perhaps suggesting that the position of the bridging ligand is intermediate to what is observed in the Hox and Hred structure [12,21,51,57,138]. The subsequent characterization of the high-resolution structures of these states by X-ray crystallographically can resolve the mode of carbon monoxide binding in these states. Recently ESEEM, ENDOR, and HYSCORE spectroscopy of native and isotopically labeled hydrogenases has been exploited to examine the electronic coupling of nitrogen atoms to the 2Fe subcluster of the H-cluster [157]. In these studies, the contribution of the nitrogen atoms of the cyanide could be assigned and confirmed the aforementioned work of Thomann, Bernardo, and Adams [140] discussed at the beginning of this section. The work also described a third coupling that was not previously characterized Met. Ions Life Sci. 2009, 6, 179–218


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and that was indicated to potentially support the presence of a secondary amine containing dithiolate (DTN) but a component of the protein environment could also account for this coupling.

4. H-CLUSTER MODEL COMPLEXES 4.1. 2Fe Subsite Model Synthesis The structures of [FeFe]-hydrogenases were received enthusiastically by the inorganic and organometallic synthetic community. This was not only due to the nature of the cluster structure, reactivity, and the prospects of a new synthetic target but also due to the fact that a sound synthetic framework for the mimetic compounds was set in place. The H-cluster is modular composed of a [4Fe-4S] subcluster bridged to the 2Fe organometallic unit with electron-donating CO and CN– ligands and also the unique non-protein dithiolate ligand. Even at the advent of the first structure determinations it was clearly perceived that the hydrogen reactivity occurred at the 2Fe unit and it was postulated that the reactivity could occur at the observed more dynamic coordination site of the distal Fe atom [20] in which either a H2O molecule, CO molecule, or a vacant site is observed in the CpI Hox state [11], CpI HoxCO state [19], and DdH Hred state [18,21] structures respectively. Therefore, the 2Fe subcluster was the focus of the first synthetic model studies [151,153,158] and much effort has been directed to replicate the unique features of the 2Fe subcluster which make hydrogen catalysis possible. Specific challenges in modeling the 2Fe subluster site include its unique geometry, depicting the role of CO/CN ligands in relation to low oxidation and spin states, low oxidation mixed valence 2Fe states, and incorporation/ role of the bridging CO ligand. Significant progress has been made in modeling the 2Fe subcluster site [151,153,158–184] and many insights have been gained not only simply regarding structural aspects of the 2Fe subcluster and its unique ligands but also how its unique features play key mechanistic and electronic roles during the hydrogen activation process. The most basic model to the complex 2Fe subcluster is diironhexacarbonyldisulfide (Figure 5A) and the synthesis of this compound can be dated back all the way to 1929 [185]. Although this compound has only the bare framework of the 2Fe subcluster, it provided the initial pathway for synthesis of more complex 2Fe subcluster models. In the 1980s, developments in organometallic synthesis demonstrated the reactivity of the sulfur and iron bonds in the diironhexacarbonyldisulfide compounds and synthesis of the all carbonyl complexes [Fe2(m-SR)2)(CO)6] with variations on the sulfur and iron ligands [186–188] were reported including the most simple dithiolate bridge Met. Ions Life Sci. 2009, 6, 179–218

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Figure 5. Overview of the general development and progression of synthetic model compounds to the 2Fe subcluster and H-cluster of [FeFe]-hydrogenases. A. Diironhexacarbonyldisulfide. B. Insertion of the PDT ligand and single cyanide substitution at each Fe atom of the hexacarbonyl compound. C. Further variation on the dithiolate ligand including insertion of nitrogen and oxygen atoms at the bridge head position. D. Development of advanced models to improve similarity and functionality of the enzyme’s 2Fe subcluster including (from left to right): mixed valance FeIFeII species with a bridging CO ligand, bridging hydride, and terminal hydride species. E. Synthesis of the basic 6Fe H-cluster framework.

(PDT) (Figure 5B) [189]. The resulting compound [Fe2(SCH2CH2CH2S) (CO)6] is often considered to be the parent model for the ensuing 2Fe subcluster models. Interestingly, this line of synthetic chemistry evolved a decade earlier from report of the first [FeFe]-hydrogenase crystal structures and identification of the CO and CN ligands at the 2Fe subcluster site by IR Met. Ions Life Sci. 2009, 6, 179–218


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studies as discussed above. Shortly after, stoichiometric cyanide substitution at each Fe center on the [Fe2(SCH2CH2CH2S)(CO)6] compound was achieved and synthesis of the [Fe2(SCH2CH2CH2S)(CO)4(CN)2]2 compound was reported by three different groups (Figure 5B) [151,153,158]. This complex already exhibited key features of the 2Fe subcluster site such has its edge shared bipyramidal geomeotry, Fe-Fe distance (2.6 A˚), and placement of an open coordination site above the distal Fe atom. Most similar to the Hred state of the DdH enzyme, the model compound exists with nonparamagnetic FeI centers. The next feature addressed by development of model complexes to the 2Fe subcluster was introduction of variations on the dithiolate ligand. Because the reported X-ray crystal structures could not distinguish between isoelectronic groups at the middle atom of the bridging ligand, it is possible that either -CH2-, -NH-, or -O- could exist at that position and synthesis of two other possible bridging ligands, (SCH2NRCH2S) R ¼ H, Me) [159] and (SCH2OCH2S) [174], was reported (Figure 5C). Looking at the case of the amine dithiolate bridge, from a mechanistic perspective the nitrogen was described as being an attractive possibility at that position as it could act as base to extract and transfer protons [131,133,190–192] at the ligandexchangeable site above the distal Fe atom during hydrogen catalysis. There is however, no crystallographic evidence to date that supports this hypothesis and it is important that model complex studies keep in mind all possibilities for the bridging ligand (PDT, DTN, DTO). Given the rapid developments of model compounds to the 2Fe subcluster site, these early studies still failed to address a stable mixed valence state of the Hox state and the bridging CO ligand present in this state due to low stability and reactivity. These two characteristics are thought to be important to H2 catalysis, especially as in the H2 catalytic cycle the H-cluster cycles between just two states (Hox and Hred). Picket and coworkers were the first able to spectrochemically characterize a mixed valence FeIFeII species with a bridging CO ligand, however due to the low stability, isolation of the compound remained elusive [169]. This work prompted the development of diferrous compounds that resemble the Hox state where an isocyanide ligand is bridging between the two Fe atoms. The diferrous compound [Fe2SCH2CH2CH2S(CNMe)7][PF6]2 closely mimics the geometry of the Hox active state and includes the presence of a bridging ligand, however, the stability of the compound can be greatly attributed to the higher oxidation state of the diiron dithiolate complex [170]. In 2007, Liu and Darensbourg reported the first isolation of a stable mixed valence FeIFeII species with a bridging CO ligand [183] and shortly after, Justice, Rauchfuss and Wilson [182] reported another similar species (Figure 5D). EPR and FTIR on the species revealed a similar rhombic EPR spectra and FTIR spectra [182,183] to the Hox DdH state and the reported Met. Ions Life Sci. 2009, 6, 179–218

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models were the first to exhibit a FeIFeII paramagnetic couple with a bridging CO ligand. Significantly, these models were a breakthrough in modeling the active form of the 2Fe subcluster, however, both included bulky strong donor ligands, IMES and dppv, in place of the CN ligand at one of the Fe atoms. These ligands were presumably used to lock the unique geometry in place with the bridging CO ligand and mixed valence FeI/FeII state and may not be effective mimics of the actual biological H-cluster ligands. In addition, at the proximal Fe, in both cases, a PMe3 ligand was used in place of the CN ligand such that these models capture the key features of the 2Fe subcluster in the active Hox state but they have not been able to include simple electron donating CN ligands and ongoing work is directed toward capture of simple FeI/FeII models containing CN ligands, a bridging CO ligand, an unsaturated Fe atom.

4.2. Advanced 2Fe Subsite and H-Cluster Model Synthesis With respect to the most recent work in this area, much emphasis is placed on modeling the hydrogenase functionality emphasizing hydride complexes and synthesis of the complete 6Fe H-cluster framework. Darensbourg et al. have reported the synthesis of 2Fe compounds containing a bridging hydride ([Fe2(SCH2CH2CH2S)(m-H)(CO)4(PMe3)2]1) (Figure 5D) [165,166]. Significantly, this compound begins to model the functionality of the 2Fe subcluster as it has the ability to bind and activate H2 at an FeII site. Although the biochemical and structural characterization have not implicated a role for a bridging hydride in the enzymes, this is the first compound to show H2 binding at an unsaturated FeII center. Terminal hydride compounds have also been synthesized and the synthesis of a 2FeI species having a bridging carbon monoxide ligand and a terminal hydride ligand has been reported (Figure 5D) [184]. In the Hox state, the bridging carbon monoxide ligand may prevent protonation of the FeFe bond such that proton binding at the distal Fe would be more favorable [193]. Development of hydride binding species is important in unraveling the mysteries still present at the 2Fe subcluster and mechanistic features of H2 catalysis. In 2005 Picket et al. reported the synthesis of an entire 6Fe H-cluster framework with a 2Fe subcluster model bridged to a [4Fe-4S] subcluster using a tridentate thiolate ligand (Figure 5E) [194]. On a similar time frame it was realized through DFT calculations that there is a signficant impact of the presence of the [4Fe-4S] cluster on the overall electronic structure of the 2Fe subcluster and likely catalysis [150,195]. This highlights the fact that the groundwork for DFT studies has been well set in place from the development of synthetic models to the 2Fe subcluster and synthetic model studies have opened opportunities for DFT studies to explore H2 oxidation and H1 Met. Ions Life Sci. 2009, 6, 179–218


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reduction at the 2Fe subcluster and the complete 6Fe cluster. The application of DFT to modeling the H-cluster during H2 catalysis will be discussed in much greater detail in Chapter 12.

5. H-CLUSTER BIOSYNTHESIS 5.1. Relevance to Prebiotic Chemistry H-cluster biosynthesis is an emerging area of research that is key to generating effective metabolic engineering of hydrogen-producing microorganisms and heterologous expression and mass production of hydrogenase enzymes for use in biomimetic and biohybrid materials in biotechnology. But as in the case of the synthesis of small molecule mimetics described in Section 4, there are links to these enzyme active sites to prebiotic chemistry [61–63] and understanding the relationship between biosynthesis, structure, and reactivity and relating these findings to the ongoing work exploring the structure and reactivity of iron-sulfur minerals and their derivatives may hold keys to life’s origins. Of course, the suggestion that iron-sulfur minerals may have had a role in prebiotic chemistry and the origin of life are not new and the potential for this has been expounded upon in recent works of Wa¨chtershaüser and others [41]. The complex iron-sulfur cluster at the active sites of nitrogenases, hydrogenases, carbon monoxide dehydrogenases, and acetyl-CoA synthase reactions have been suggested to be among the first cofactors since these enzymes catalyze interconversions of small molecules that could have been important in increasing the pool of reactants to participate in basic condensation reactions important for forming the building blocks of life. These enzymes, all having deeply rooted lineages in a ‘‘metabolism first’’ model for the origin of life, may represent highly evolved derivatives of iron-sulfur minerals which supported the reactivity to make the transition from the non-living to the living world. These ideas are dependent on a mechanism to generate a variety or combinatorial allotment of modified iron-sulfur mineral catalysts tuned to specific reactions equivalent to enzymes in a metabolic pathway. We anticipate that reduced iron-sulfur minerals would have been plentiful on the early earth and the means to generate a variety of structures of modified iron-sulfur minerals exists today in environments that many feel are closest to mimics of early earth’s environments. Examples of these environments are the hydrothermal vents in the deep oceans or the effluent streams leading from thermal springs like those found in Yellowstone National Park. These environments represent highly mineral-rich gradients and those rich in iron and sulfur are not difficult to find today. With large Met. Ions Life Sci. 2009, 6, 179–218

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gradients in temperature, pH, gas solubility, mineral solubility, etc. it is not difficult to imagine the introduction of carbon monoxide or heteroatoms such as Mo or Ni to tune various types of reactivity. A complete understanding of the structure and reactivity of these complex iron-sulfur enzymes together with a full understanding of the chemical reactions that bring about the biosynthesis of the prosthetic groups will provide the basis for a better understanding of the potential for generating modified iron-sulfur minerals that resemble these clusters on the early earth.

5.2. Identification of Genes Until only very recently essentially nothing was known about [FeFe]hydrogenase H-cluster biosynthesis. One of the reasons for this might be that there has been limited success in terms of genetically manipulating organisms that harbor [FeFe]-hydrogenases. For the nitrogenase system, genetic studies were key in the identification of genes other than the nitrogenase structural genes nifH, nifD, and nifK required for the process of nitrogen fixation (see reviews [196,197]). These studies provided significant insights into understanding the regulation of nitrogen fixation, the linkages between nitrogen fixation and energy metabolism, and iron-sulfur cluster biosynthesis. Gene products were initially implicated in being involved in iron-sulfur cluster biosynthesis through the analysis of nitrogenases produced in various genetic backgrounds in which nitrogen fixation specific gene products had either insertion or deletion mutations. In these genetic backgrounds it was found that a number of mutants produced nitrogenases that had a complement of iron-sulfur clusters that differed from the native enzyme. From these studies a number of gene products were specifically implicated in being involved in nitrogenase iron-molybdenum cofactor biosynthesis. This approach and the analysis of nitrogenase mutants later proved to be the driving force in some of the first breakthrough studies that provided insights into general mechanisms of iron-sulfur cluster biosynthesis. Some interesting insights have been made into the active site cluster biosynthesis of [NiFe]-hydrogenases including the source of cyanide ligands (discussed in detail in Chapter 5) [198–202]. For [FeFe]-hydrogenase H-cluster biosynthesis, it was not until 2004 when an approach similar to that described for the identification of nitrogen fixation gene products involved in cluster biosynthesis was applied [203]. In this work, several strains of Chlamydomonas reinhardtii produced by transposon mutagenesis that were unable to produce hydrogen were identified. These strains were analyzed with regard to what specific genes were interrupted to eliminate hydrogen production. Several gene disruptions were identified that were not Met. Ions Life Sci. 2009, 6, 179–218


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localized to either of the two [FeFe]-hydrogenase structural genes present in C. reinhardtii including within a gene cluster encoding two novel members of the S-adenosylmethionine dependent (radical-SAM) enzymes. These gene products termed hydEF and hydG were not found to be clustered in the C. reinhardtii genome with either of the [FeFe]-hydrogenase structural genes hydA, however, it was found that the expression of the [FeFe]-hydrogenase in a background of co-expressed HydEF and HydG heterologously in E. coli resulted in the synthesis of active [FeFe]-hydrogenase [203]. Although only a fraction of the anticipated activity (B20%) was obtained, expression of the [FeFe]-hydrogenases in E. coli without co-expression of the identified accessory enzymes does not yield any active enzyme. The results are highly significant and the ability to synthesize an active enzyme having such a unique active site cluster will be tremendously valuable in defining the biochemistry of H-cluster biosynthesis. For nitrogenase, the analogous feat of expression of active enzyme in an E. coli host has not been achieved.

5.3. Gene Clusters and Operons These studies provide a framework for thinking about how the H-cluster is synthesized. A closer look at the deduced amino acid sequences of hydEF and hydG and their homologs in other organisms provides additional insights. All other organisms that are known to harbor active [FeFe]hydrogenases have homologs of the C. reinhardtii genes, however, in other organisms hydE and hydF occur as separate genes. In addition, although in many cases the genes hydE, hydF, and hydG and the structural gene(s), do not occur in gene clusters and are distributed throughout organisms essentially indiscriminately, there are several examples in which the genes do occur in clusters (Figure 6). In a number of cases these clusters contain genes, in addition to hydE, hydF, and hydG, and the structural gene(s) however, these are the only genes yet to be determined to be common in all organisms. Interestingly the aal gene product ammonia aspartate lyase is present in a number of gene clusters but is not universal in its occurrence with hydrogen genes. The occurrence of ammonia aspartate lyase may implicate a link between biosynthesis and central carbohydrate or amino acid metabolism. Thus it is interesting to contemplate about [FeFe]-hydrogenase in the context of only these products of the three genes hydE, hydF, and hydG. One thing that is clear in comparing the sequences and properties of genes involved in the [FeFe]- and [NiFe]-hydrogenases (Chapter 5) is that the biochemistry of cluster synthesis and hydrogenase maturation in the two systems is very different. Although there is good evidence that the source of cyanide ligands in the [NiFe]-hydrogenases is carbamoyl phosphate Met. Ions Life Sci. 2009, 6, 179–218

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Figure 6. Gene clusters of various [FeFe]-hydrogenases showing the observed genomic arrangement of [FeFe]-hydrogenase maturation gene products HydE, HydF, and HydG along with the [FeFe]-hydrogenase structure encoding gene product HydA. Also present in some operons is AspA ammonia aspartate lyase encoded by the aal gene.

[198,199], this similarity is not found to occur with [FeFe]-hydrogenase genes and thus, cyanide is most likely to be derived from another source. Some may consider the enzyme carbon monoxide dehydrogenase which catalyzes reversible carbon monoxide oxidation to be a potential mechanism for deriving carbon monoxide ligands in both [FeFe]- and [NiFe]-hydrogenases, however, this possibility can be dismissed. The processes of carbon monoxide oxidation and anaerobic carbon dioxide fixation occur much less frequently in biology than hydrogen metabolism and thus, many if not most hydrogenases occur in organisms that do not possess a carbon monoxide dehydrogenase. In addition, the only cases in which the genes encoding carbon monoxide dehydrogenase occur with hydrogenases are in cases where carbon monoxide oxidation is metabolically coupled to hydrogen metabolism as in the case of the purple non-sulfur bacteria [204], making the hypothesis of carbon monoxide derivation for active site formation necessitated by an active hydrogenase untenable.

5.4. Radical-S-Adenosylmethionine Chemistry and Potential Biological/Biochemical Sources We have thought long and hard about the potential of only three gene products in H-cluster biosynthesis. This would be in stark contrast to the nitrogenase FeMo cofactor biosynthesis in which approximately ten gene products can be genetically and biochemically linked to the process [196,197]. However, making several assumptions based on the observations associated with FeMo cofactor biosynthesis and taking into account the clear implications concerning the potential function of the H-cluster Met. Ions Life Sci. 2009, 6, 179–218


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accessory gene annotation and preliminary biochemical characterization has led to the ability to suggest possible rationals for H-cluster formation. The observation that the FeS cluster containing proteins HydE and HydG are clearly radical-SAM enzymes as by the observance that they both have the C-X3-C-X2-C radical-SAM signature motif (Figure 7) [203,205] and that

Figure 7. A. Portion of sequence alignment of radical-SAM [FeFe]-hydrogenase proteins HydE and HydG relative to the known functional radical-SAM proteins biotin synthase (BioB), lipoate synthase (LipA), pyruvate formate lyase activating enzyme (PFL-AE), lysine 2,3-aminomutase (LAM), and thiazole synthase (ThiH). The signature radical-SAM motif C-X3-C-X2-C is highlighted red and conserved and similar residues are highlighted light blue and green, respectively. B. Reactions catalyzed by radical-SAM proteins. BioB and and LipA catalyze hydrogen atom abstraction/sulfur insertion reactions and PFL-AE, LAM, and ThiH catalyze various reactions via the formation of an amino acid radical. Met. Ions Life Sci. 2009, 6, 179–218

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HydF protein is a GTPase [203,206,207] gives a solid foundation to think about H-cluster biosynthesis. In fact, radical-SAM chemistry alone clues for potential mechanism of H-cluster biosynthesis. In 2007, we published an article and detailed a potential hypothetical mechanism for H-cluster biosynthesis based on what is known concerning FeMo cofactor biosynthesis together with observations and considerations of the H-cluster structure and the involvement of radical-SAM based biochemistry [208]. We had made several assumptions to frame our hypothetical mechanism. One of the most important assumptions was that the maturation machinery was directed only at the 2Fe subcluster of the H-cluster. We made this assumption for several reasons, including (1) the limited number of gene products implicated in H-cluster biosynthesis [203], (2) the ability of organisms processing [FeFe]-hydrogenase to synthesize [4Fe-4S] clusters [209], and (3) the observation that the only covalent linkage between the two subclusters is provided by the protein [11,18,21]. The other important assumption we made was that the synthesis of the 2Fe subcluster does not occur on the hydrogenase and occurs on a protein scaffold in analogy to nitrogenase FeMo cofactor biosynthesis [210] and thus, we anticipate that the [FeFe]-hydrogenase produced in a background devoid of the maturation genes should be produced in a form that is capable of being activated upon the insertion of the 2Fe subcluster. With this foundation we then began to consider the involvement of radical-SAM chemistry and specifically tandem radical-SAM enzymes catalyzing sequential steps in the process which would include the synthesis and placement of the non-protein ligands and production of the 2Fe subcluster. In the context of precedented radical-SAM chemistry, one connection that appears to be intuitive is that sulfur insertion chemistry similar to that observed in BioB [211] and LipA [212] in the synthesis of biotin and lipoic acid is likely invoked in the synthesis of the dithiolate ligand (Figure 7). In these reactions it has been shown that the source of sulfur for these cofactors originates from a second conserved FeS cluster unique from the SAM-binding [4Fe-4S] cluster [213–217]. For this reaction, it is not intuitive what might be the substrate for the sulfur insertion. It has been suggested that perhaps propane or dimethylamine could serve as substrates resulting in a PDT or DTN ligand, respectively, but these substrates are hard to rationalize in the context of microbial metabolism. Since only there are only three gene products and there are no clear conserved gene products that would be involved in providing the substrates for either of the radical-SAM enzymes, it may be rational to suggest that the precursors come from the central metabolism and in the case of [FeFe]-hydrogenase this would be specific to anaerobic metabolism. Although this perhaps narrows the possibilities, there are still a large number of compounds that could serve as precursors and since the composition of Met. Ions Life Sci. 2009, 6, 179–218


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the dithiolate has yet to be determined, this makes rationalizing potential substrates even more difficult. We have therefore kept our thoughts on this step of cluster biosynthesis fairly generalized with the notion that the reaction, whatever the precursor, resembles the reactivity observed for LipA in that a sulfur insertion occurs by the hydrogen atom abstraction at two positions of an organic substrate [215] and the source of the thiolate sulfurs are sulfides from an FeS cluster. In contrast to the LipA reaction in which the FeS cluster is destroyed in the process of lipoic acid synthesis, we envision a reaction in which a [2Fe-2S] cluster is converted to a dithiolatecoordinated 2Fe cluster that remains intact. If we think about this reaction as the first step of H-cluster biosynthesis and we examine this from the perspective of the reactivity of a hypothetical [2Fe-2S] precursor, the reactive sulfides are converted to less reactive thiolates allowing us to think about a second step of the biosynthetic pathway in which the Fe ions are the focus of the reactivity. Additional ideas can be gleaned from precedented radical-SAM chemistry. One set of reactions or themes that we found particularly attractive are reactions involving amino acid radicals as intermediates. The reactions catalyzed by pyruvate formate lyase activating enzyme [218], lysine amino mutase [219], and the thiH gene product [220,221] involved in synthesis of the thiazole ring of thiamin all occur via the formation of amino acid radicals (Figure 7). The observation of multiple examples of radical-SAM enzymes that utilize amino acids as substrates and catalyze bioconversions of amino acids involving the formation of amino acid radicals makes it attractive to think of an amino acid substrate and an amino acid radical intermediate as a source of the carbon monoxide and cyanide ligands in the H-cluster. There are some attractive features with regard to the potential utilization of an amino acid as a substrate for carbon monoxide and cyanide formation, probably the most significant being that an amino acid substrate and the decomposition of an amino acid at a iron-sulfur cluster as a means for forming these ligands represents a mechanism in which both carbon monoxide and cyanide ligands can be formed and deposited on the metal center in a concerted manner from a fixed source such that free carbon monoxide and cyanide do not freely diffuse through cells. Considering the availability of substrates that could be involved in a reaction forming carbon monoxide and cyanide in a concerted manner it is attractive to think about the decomposition of an amino acid. In our hypothesis we suggested glycine as a potential substrate for a radical-SAM enzyme yielding upon decomposition carbon monoxide, cyanide, and water, essentially the ligand set observed at the distal Fe of the 2Fe subcluster [208]. Glycine is an interesting potential substrate from the perspective of availability since glycine is required for nucleotide metabolism and thus, should be available in all organisms. A computational analysis of the Met. Ions Life Sci. 2009, 6, 179–218

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viability of such a reaction was analyzed using a hypothetical in-silico generated simple thiolate compound and yielded an overall thermodynamically favorable reaction with an activation barrier for the formation of the glycine radical that binds and decomposes at the site. The barrier can be overcome by the activity of a radical-SAM enzyme perhaps lending credence to this overall concept. These ideas are interesting but not founded in experimental data and only provide the basis for experimental design to prove or disprove them. Another interesting perspective of the idea of an amino acid decomposition as a component of H-cluster biosynthesis returns us to the idea of a link between iron-sulfur enzymes and prebiotic chemistry. If we consider the reverse of the reaction described above we have the condensation of carbon monoxide, and water to form an amino acid. In essence, the formation of a building block from small molecule precursors is conceptually similar to the early experiments of Miller and Urey [222] but adapted in the spirit of the current ideas of Wa¨chtershaüser in clearly identifying a central role for iron-sulfur compounds in early Haedian and prebiotic reactions [41,43,223].

6. FUTURE DIRECTIONS The ability to examine the above ideas concerning the formation of the nonprotein ligands (dithiolate, carbon monoxide, and cyanide) and screen for the actual substrates for radical-SAM enzymes (HydE and HydG) is dependent on a functional assay for their activities. A recent advancement in this regard is the development of an in vitro system in which it has been shown that a hydrogenase structural gene product (HydA) from Clostridium or Chlamydomonas expressed in an E. coli host in the absence of any accessory enzymes (without HydE, HydF, or HydG) is capable of being converted into an active hydrogenase by the addition of E. coli cell extracts in which all three accessory enzymes are expressed in concert [224]. The results of this work suggest that the bulk of cluster biosynthesis occurs at a site other than the structural enzyme implicating a potential role of some scaffold in the process. Our recent results implicate a role of HydF in this process and one could envision that a precursor of the intact H-cluster could be synthesized on HydF in a manner involving sequential radical-SAM catalyzed reactions steps as described above (Figure 8). The in vitro system is just a small step in unraveling the complex process of H-cluster biosynthesis but an important step in defining a viable biochemical in vitro assay to design future experiments probing the function of the radical-SAM enzymes specifically. Met. Ions Life Sci. 2009, 6, 179–218


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Figure 8. Schematic representation depicting the proposed chemical steps of H-cluster biosynthesis involving radical-SAM enzyme activities. The 2Fe subcluster assemblies and intermediates are represented as ball-and-stick models with carbon atoms in gray, oxygen in red, nitrogen in blue, sulfur in orange, and iron in rust brown. The central atom of the dithiolate linkage and the unknown iron ligands are represented in magenta. The ribbon representation of the overall structure of the Fe-only hydrogenase is from Clostridium pasteurianum (CpI).

ACKNOWLEDGMENTS Hydrogenase research in this lab is supported by grants from the AirForce Office of Scientific Research (FA9550-05-01-0365), the Department of Energy (DE-FC36-06-GO86060 and DE-FG02-07ER46477), and the NASA Astrobiology Biogeocatalysis Center of Montana State University Funded by NASA (NNA08CN85A). Portions of this research were carried out at the Standford Synchrotron Radiation Laboratory, a national user facility operated by Stanford University on behalf of the U.S. Department of Energy, Office of Basic Energy Sciences. The SSRL Met. Ions Life Sci. 2009, 6, 179–218

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Structural Molecular Biology Program is supported by the Department of Energy, Office of Biological and Environmental Research and by the National Institutes of Health, National Center for Research Resources, Biomedical Technology Program, and the National Institute of General Medical Sciences. Special thanks to David Mulder for assistance in preparation of this chapter.

ABBREVIATIONS AND DEFINITIONS BioB CpI Crl DdH DFT dppv DTN DvH ENDOR EPR ESEEM FTIR HYSCORE IMES IR LAM LipA NADH PDT PFL-AE SAM Tma ThiH

biotin synthase Clostridium pasteurianum [FeFe]-hydrogenase Chlamydomonas reinhardtii [FeFe]-hydrogenase Desulfovibrio desulfuricans [FeFe]-hydrogenase density functional theory cis-1,2-C2H2(PPh2)2 dithiolate Desulfovibrio vulgaris Hildenborough [FeFe]-hydrogenase electron nuclear double resonance spectroscopy electron paramagnetic resonance spectroscopy electron spin echo envelope modulation spectroscopy Fourier transform infrared spectroscopy hyperfine sublevel correlation spectroscopy 1,3-bis(2,4,6-trimethylphenyl)imidazol-2-ylidene infrared lysine 2,3-aminomutase lipoate synthase nicotinamide adenine dinucleotide, reduced propanedithiolate pyruvate formate lyase activating enzyme S-adenosylmethionine Thermotoga maritima [FeFe]-hydrogenase thiazole synthase

REFERENCES 1. R. P. Happe, W. Roseboom, A. J. Pierik, S. P. Albracht and K. A. Bagley, Nature, 1997, 385, 126. 2. M. W. Adams, Biochim. Biophys. Acta, 1990, 1020, 115–145. 3. A. E. Przybyla, J. Robbins, N. Menon and H. D. Peck Jr., FEMS Microbiol. Rev., 1992, 8, 109–135.

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7 Carbon Monoxide as Intrinsic Ligand to Iron in the Active Site of [Fe]-Hydrogenase Seigo Shima,a Rudolf K. Thauer, a and Ulrich Ermler b a

Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Strasse, D-35043 Marburg, Germany b Max Planck Institute for Biophysics, Max-von-Laue-Strasse 3, D-60438 Frankfurt/Main, Germany

ABSTRACT 1. INTRODUCTION 2. PHYSIOLOGY 3. THE IRON GUANYLYLPYRIDINOL COFACTOR IN THE ENZYME-FREE STATE 3.1. Isolation of the Iron Guanylylpyridinol Cofactor and Activity Assay 3.2. Proposed Structure of the Iron Guanylylpyridinol Cofactor 3.3. Electronic and Magnetic State of the Iron 3.4. Stability of the Iron Guanylylpyridinol Cofactor 4. STRUCTURE OF [Fe]-HYDROGENASE WITH AND WITHOUT THE IRON GUANYLYLPYRIDINOL COFACTOR BOUND 4.1. Apo- and Holoenzyme Production 4.2. Crystal Structure Determination 4.3. Protein Fold 4.4. The Binding Sites for the Iron Guanylylpyridinol Cofactor and for Methenyl-H4MPT1


220 220 222 223 223 224 225 225

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5. LIGANDS TO IRON IN THE ACTIVE SITE OF [Fe]HYDROGENASE 5.1. Intrinsic Ligands 5.2. Carbon Monoxide and Cyanide as Extrinsic Ligands 5.3. Inactivation of [Fe]-Hydrogenase by O 2 and Cu(I) 6. PROPOSED CATALYTIC MECHANISMS 6.1. H2 Binds First to the Carbocationic C14a of Methenyl-H4MPT1 6.2. H2 Binds First to the Active Site Iron Carbonyl Complex 7. CONCLUDING REMARKS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

231 231 232 234 235 235 236 237 237 237 238

ABSTRACT: Structural and spectroscopic studies on [Fe]-hydrogenase revealed an active site mononuclear low spin iron coordinated by the Cys176 sulfur, two CO, and the sp2 hybridized nitrogen of a 2-pyridinol compound with back bonding properties similar to those of cyanide. Thus, [Fe]-hydrogenases are endowed with an iron-ligation pattern related to that found in the active site of [NiFe]- and [FeFe]-hydrogenases although the three hydrogenases and the enzymes involved in their posttranslational maturation have evolved independently and although CO and cyanide ligands are not found in any other metallo-enzymes. Obviously, low-spin iron complexed with thiolate(s), CO, and cyanide or a cyanide functional analogue plays an essential role in H2 activation. KEYWORDS: carbon monoxide cyanide hydrogen activation hydrogenase iron carbonyl iron guanylylpyridinol cofactor methanogenic archaea

1. INTRODUCTION [Fe]-hydrogenase was discovered in 1990 and according to its systematic name was referred to as H2-forming methylenetetrahydromethanopterin dehydrogenase, abbreviated Hmd, which is also the abbreviation for the encoding gene (hmd) [1,2]. The enzyme was found as a homodimer with a molecular mass of ca. 80 kDa that contains 2 iron per mol homodimer but no iron-sulfur clusters [1,2]. The iron was first thought to be non-functional, mainly because of its redox inactivity, and therefore the enzyme was addressed as metal-free hydrogenase [3,4] in contrast to the [NiFe]- and [FeFe]-hydrogenases. It had been overlooked that the enzyme is reversibly inhibited by CO and by cyanide, albeit only at relatively high concentrations. Especially CO inhibition indicates an involvement of the iron in catalysis. It took until 2004 to demonstrate unambiguously that the iron is essentially required for the catalytic activity of the enzyme [5]. After another two years the iron center was definitively identified as mononuclear [6] and not Met. Ions Life Sci. 2009, 6, 219–240

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dinuclear as in [NiFe]-hydrogenases (see Chapter 5 of this volume) and [FeFe]-hydrogenases (Chapter 6) and the enzyme was finally named as [Fe]-hydrogenase. Whereas [NiFe]-hydrogenases are found in many archaea and bacteria and [FeFe]-hydrogenases in bacteria and lower eucarya, the occurrence of [Fe]-hydrogenase appears to be restricted to methanogenic archaea growing on H2 and CO2 as sole energy source. The reason for this is that [Fe]hydrogenase catalyzes one specific reaction in their energy metabolism [7] namely the reduction of methenyltetrahydromethanopterin (methenylH4MPT1) with H2 to methylenetetrahydromethanopterin (methyleneH4MPT) (reaction 1) (Figure 1) [8,9]. H2 þ methenyl-H4 MPTþ Ð methylene-H4 MPT þ Hþ DG0 0 ¼ 5:5 kJ=mol

ð1Þ

[Fe]-hydrogenase contains tightly bound an iron guanylylpyridinol cofactor (FeGP cofactor). The iron is ligated to the guanylylpyridinol molecule, 2 CO, and the sulfur of Cys176 which covalently links the FeGP cofactor to the protein [6]. This ligation pattern is reminescent to that of [FeFe]- and [NiFe]-hydrogenases which also contain an iron complexed by CO and by cyanide. The pyridinol moiety of FeGP might be a functional analogue to cyanide (see below). Interestingly, neither [Fe]-hydrogenases, [FeFe]hydrogenases and [NiFe]-hydrogenases nor the enzymes involved in their posttranslational maturation, as far as known, are phylogenetically related and intrinsic CO and cyanide ligands are not found in any other metalloenzyme [10]. This finding indicates that the H2 activation sites in the three enzymes have evolved convergently [11] and attributes to them a unique function in H2 activation.

Figure 1. Reaction catalyzed by [Fe]-hydrogenase (Hmd). A hydride is transferred from H2 to C14a of methenyl-H4MPT1 from the pro-R side yielding methyleneH4MPT and a proton [8,9]. The methenyl C14a has carbocation character and is therefore an excellent hydride acceptor [32–34]. H4MPT, tetrahydromethanopterin. Met. Ions Life Sci. 2009, 6, 219–240

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The definition of [Fe]-hydrogenase as ‘‘hydrogenase’’ can be questioned. The pros are that the enzyme catalyzes a reversible reaction with H2 as substrate and that in H2 activation it involves an active site iron carbonyl as do [NiFe]- and [FeFe]-hydrogenases (see Chapters 5 and 6). The con is that [Fe]hydrogenase exhibits a ternary complex catalytic mechanism [12] whereas [NiFe]- and [FeFe]-hydrogenases show ping-pong catalytic mechanisms. Thus, [Fe]-hydrogenase does not react with H2 in the absence of its substrate methenyl-H4MPT1 [13] whereas [NiFe]- and [FeFe]-hydrogenases are reduced by H2 in the absence of an external hydride or electron acceptor. This difference is reflected in the finding that [Fe]-hydrogenase catalyzes a single and double exchange of H2 with protons of water only in the presence of methenylH4MPT1 [12–16] whereas [NiFe]- and [FeFe]-hydrogenases do this per se. The literature on [Fe]-hydrogenase has recently been reviewed by the authors [17]. This chapter will therefore concentrate on the FeGP cofactor, in particular, on the ligation pattern of the iron.

2. PHYSIOLOGY In methanogens reaction (1) is also catalyzed by two other enzymes working together, namely by F420-reducing [NiFe]-hydrogenase (Frh) and F420-dependent methylenetetrahydromethanopterin dehydrogenase (Mtd), which catalyze reactions (2) and (3), respectively [18]. H2 þ F420 Ð F420 H2

DGo 0 ¼ 11 kJ=mol

F420 H2 þ methenyl-H4 MPTþ Ð F420 þ methylene-H4 MPT þ Hþ DG0 0 ¼ þ5:5 kJ=mol

ð2Þ ð3Þ

This explains why not all methanogens have to contain [Fe]-hydrogenase. The enzyme is absent in all investigated members of the Methanosarcinales, in most members of the Methanomicrobiales, and some members of the Methanobacteriales. The methanogens, that contain [Fe]-hydrogenase, are adapted to niches, where the H2 partial pressure is relatively high and the nickel concentration can be low. This is deduced from the finding that the apparent Km for H2 of [Fe]-hydrogenase is ten times higher than that of the F420-reducing [NiFe]-hydrogenases and that the [Fe]-hydrogenase is overproduced in the methanogens under nickel limiting growth conditions, under which the F420-reducing [NiFe]-hydrogenases are not synthesized [18,19]. [Fe]-hydrogenase has been found in methanogens thriving in hot vents (Methanopyrus Met. Ions Life Sci. 2009, 6, 219–240


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kandleri [20] and Methanocaldococcus jannaschii [21]), in salt marshes (Methanococcus maripaludis [22]), in anaerobic sewage digestion plants (Methanothermobacter thermoautotrophicus [23] and M. marburgensis [1]) and the human intestinal tract (Methanobrevibacter smithii [24]). After growth of M. marburgensis under nickel limiting conditions the cell contains [Fe]-hydrogenase at a concentration of nearly 5% of its soluble proteins, from which the enzyme can be purified to homogeneity in high yields [18]. All methanogens with [Fe]-hydrogenase were found to harbour a gene nikR, a homolog of which encodes for a protein NikR that senses the intracellular nickel concentration and binds to the promoter region of the nickel importer genes nikABCDE and represses their transcription when containing nickel bound [25]. In E. coli NikR binds specifically to a palindrome sequence (CTATGA-N16-TCATAG) [26]. A similar palindrome sequence is also found in the promoter region of the hmd gene in M. marburgensis (GTACTAC-N14-GTATTAC), M. maripaludis (GTATTA-N15ATATTAC), and M. jannaschii (ATATTAC-N14-ATATTAC [25,27].

3. THE IRON GUANYLYLPYRIDINOL COFACTOR IN THE ENZYME-FREE STATE Most of what is known about the cofactor has been worked out with the cofactor of [Fe]-hydrogenase from M. marburgensis. This thermophilic methanogen is easy to grow under nickel limiting conditions in 100 g (wet mass) amounts, from which approximately 100 mg of enzyme are purified to homogeneity in a yield of 50%.

3.1. Isolation of the Iron Guanylylpyridinol Cofactor and Activity Assay From 100 mg of [Fe]-hydrogenase (1.25 mmol homodimer) approximately 2 mmol cofactor are obtained in a procedure, in which the enzyme solution is supplemented with methanol (60% final concentration), ammonia (600 mM) and mercaptoethanol (1 mM). Mercaptoethanol or other thiol compounds thereby displace the cysteine sulfur ligand to iron through which the FeGP cofactor is covalently bound to the protein (reaction 4). After incubation for 16 hours at 4 1C, 1 M NaCl in 50 mM Tris/HCl pH 8.0 is added to a final concentration of 0.2 M and the precipitated material removed by centrifugation and the cofactor purified by HPLC [28]. Active enzyme (Hmd holoenzyme) is regenerated upon addition of the extracted cofactor to refolded or Met. Ions Life Sci. 2009, 6, 219–240

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heterologously produced Hmd apoenzyme (reaction 5) providing an activity assay for the FeGP cofactor [4]. ðin 60% mehanolÞ

½Fe-hydrogenase þ mercaptoethanol ! unfolded Hmd apoenzyme þ FeGP cofactor folded Hmd apoenzyme þ FeGP cofactor ! ½Fe-hydrogenase þ mercaptoethanol

ð4Þ

ð5Þ

3.2. Proposed Structure of the Iron Guanylylpyridinol Cofactor The structure of the protein-free FeGP cofactor is not yet know but can be deduced from the decomposition products after irradiation of the cofactor with white light [5], from the FTIR spectrum [29], and from the XAS spectra, which indicates that the iron in the cofactor is complexed by 2 CO, 1 sulfur and two O/N ligands [6], and from the crystal structure of the [Fe]-hydrogenase holoenzyme [11]. The S ligand is most probably provided by mercaptoethanol required to detach the cofactor from the protein (reaction 4), to which the cofactor is covalently bound via a cysteine sulfur-iron bond. One of the two O/N ligands is provided by the pyridone nitrogen. The other O/N ligand is most likely an oxygen of the 2-carboxymethyl group of the pyridinol as deduced from the finding that during MALDI TOF MS of the FeGP cofactor the carboxyl group is lost whereas during MS of the guanylylpyridone it is not [28]. The proposed structure is shown in Figure 2.

Figure 2. Proposed structure of the protein-free FeGP cofactor. The structure of the iron-free guanylylpyridone was elucidated by NMR and mass spectrometry [28]. The structure of the iron site is deduced from IR spectra [29], Mo¨ssbauer spectra [30], and XA spectra [6]. The results are also consistent with a structure in which the carboxyl of the carboxymethyl group of the pyridinol forms a Fe-CO-CH2- rather than a Fe-O-CO-CH2- bond and in which one of the two CO are bound to the Fe opposite to the pyridinol nitrogen. Met. Ions Life Sci. 2009, 6, 219–240


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Does the [Fe]-hydrogenase cofactor have the same structure in all methanogens? Probably yes. The cofactor can be extracted from cell extracts and its mass subsequently determined by MALDI TOF MS. All methanogens investigated that contain [Fe]-hydrogenase also possess a cofactor with a guanylylpyridone ligand. The extracted cofactor could also always reconstitute active enzyme from apoenzyme. What is known about FeGP cofactor biosynthesis? Only that one of the two methyl groups attached to the pyridine ring is derived from methionine as revealed by mass spectrometry of the cofactor isolated from cells grown in the presence of [methyl-13C]-methionine (unpublished results).

3.3. Electronic and Magnetic State of the Iron Mo¨ssbauer spectroscopy of the 57Fe-labelled FeGP cofactor in the absence and presence of an external magnetic field revealed that in the protein-free and in the protein-bound state the iron in the cofactor is in a low oxidation and spin state, either low spin Fe(0) or low spin Fe(II) [30]. Therefore, the oxidation state is not yet clear. In favor of Fe(0) is that in the FeGP cofactor it appears to be pentacoordinated which is characteristic for iron in Fe(0) complexes rather than for iron in low spin Fe(II) complexes, in which the iron is generally hexacoordinated. Note however, that with the methods employed (X-ray diffraction and XAS) a proton or H2 as sixth ligand cannot be seen. In agreement with Fe(0) is also the finding that the FeGP cofactor is not very stable in water and that the stability decreases with increasing proton concentrations. Fe(0) complexes generally slowly react with protons yielding Fe(II) and H2. However, many other properties of the FeGP cofactor are in favor of Fe(II): (i) the cofactor is very light-sensitive which Fe(0) carbonyl complexes are generally not, although also Fe(0) carbonyls decay upon irradiation with light. (ii) The cofactor does not autoxidize in the presence of O2, which Fe(0) complexes generally do. (iii) The two intrinsic CO bound to the iron in the cofactor do not exchange with extrinsic 13CO as revealed by FTIR spectroscopy [5]. Rapid exchange of extrinsic with intrinsic CO is a property characteristic for Fe(0) CO complexes. And finally (iv) the cofactor does not give rise to the formation of H2 upon acidification, which Fe(0) complexes generally do (unpublished results).

3.4. Stability of the Iron Guanylylpyridinol Cofactor The protein-free FeGP cofactor is very labile and is therefore very difficult to purify to homogeneity. Solutions have to be kept in the dark, at low Met. Ions Life Sci. 2009, 6, 219–240

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temperatures and at 4pH 9, and have to be supplemented with mercaptoethanol or other thiol compounds for the cofactor to remain intact. But even under these conditions the FeGP cofactor has a half life of only a few days. The solutions have to be frozen ato–80 1C to fully retain cofactor activity. The FeGP cofactor absorbs light in the UV-A/blue region (350–462 nm tested) (Figure 3A). Upon irradiation with UV-A/blue light at 4 1C, the color is irreversibly bleached indicating that the cofactor has decomposed [5]. Decomposition products are an iron, a guanylylpyridone (GP) [28] (Figure 3), two CO, and most probably one mercaptoethanol (reaction 6): UV-A FeGP cofactor ! Fe þ GP þ 2 CO þ mercaptoethanol

ð6Þ

The light sensitivity and the decomposition products substantiate that the cofactor is an iron carbonyl which can also be deduced from the FTIR

Figure 3. Spectra of the protein-free FeGP cofactor. (A) UV-visible spectrum of the FeGP cofactor: black line (e360E2 mM1 cm1; e300E5.5 mM1 cm1) before irradiation with white light; blue line, after 30 min irradiation; red line (e300E9 mM1 cm1) after 60 min irradiation [5]. (B) Infrared absorption in the region between 2100 and 1800 cm1. Reproduced from Lyon et al. [29] with permission of the American Chemical Society, copyright (2004). Met. Ions Life Sci. 2009, 6, 219–240


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spectrum (Figure 3B) showing two almost equally intense bands at wave numbers 2032 and 1973, respectively, characteristic for two CO bound to iron in an angle of 901 [5]. In the absence of mercaptoethanol the cofactor decomposes within minutes already at room temperature in the dark. The stability increases with the mercaptoethanol concentration up to 1 mM indicating that the thiol compound probably competes with water in binding to the iron and that the complex with a water ligand is less stable than the one with a sulfur ligand which would not be too unusual for iron carbonyl complexes. Interesting in this respect is the facile exchange of the sulfur ligand. Upon addition of the FeGP cofactor to apoenzyme (reaction 5) it takes only a few seconds until the reconstitution to the active enzyme is completed. At mercaptoethanol concentrations above 10 mM the rate of FeGP cofactor decomposition increases again. An explanation for the decrease in stability might be that at higher mercaptoethanol concentrations two thiol groups bind to the iron in the FeGP cofactor and thereby destabilize the complex.

4. STRUCTURE OF [Fe]-HYDROGENASE WITH AND WITHOUT THE IRON GUANYLYLPYRIDINOL COFACTOR BOUND Although [Fe]-hydrogenase from M. marburgensis is easy to purify in relatively large amount it was not yet possible to obtain a crystal structure of the enzyme from this organism. Only perfectly merohedrally twinned crystals were obtained that did not allow phase determination. Purification of the enzyme from other methanogens proved almost impossible because their cells do not grow well under the nickel-limiting growth conditions required to induce [Fe]-hydrogenase biosynthesis. But is was possible to heterologously produce the [Fe]-hydrogenase apoenzyme from two methanogens in E. coli and reconstitute the holoenzyme with FeGP cofactor isolated from the enzyme from M. marburgensis.

4.1. Apo- and Holoenzyme Production Unfortunately, the heterologously produced apoprotein from most methanogens is recovered in the inclusion body fraction, from which the native apoprotein is difficult to refold. Until now only the heterologous expression of the hmd gene from Methanocaldococcus jannaschii and Methanopyrus kandleri yielded fully soluble apoprotein, which was correctly folded as Met. Ions Life Sci. 2009, 6, 219–240

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deduced from the successful reconstitution of the fully active holoenzyme upon addition of the FeGP cofactor [4]. Purification of the recombinant apoprotein from E. coli was essentially performed by heating the soluble cell fraction to a temperature of 70 1C (M. jannaschii) or 80 1C (M. kandleri) and by hydrophobic interaction chromatography with a phenyl-sepharose column. The yields were ca. 40 mg/10 g wet cells, which – after reconstitution to the holoenzyme – was sufficient for spectroscopic and structural studies. The apoenzyme of M. jannaschii requires the presence of dithiothreitol to remain in solution and to retain its ability to reconstitute active enzyme with the FeGP cofactor. In the structure of the apoenzyme from M. jannaschii there are two cysteine residues, those of Cys66 and Cys118, positioned such that they can form a disulfide bridge. The two cysteines are not conserved in the M. kandleri apoenzyme, which remains stable in the absence of dithiothreitol.

4.2. Crystal Structure Determination Crystals suitable for X-ray structure analysis were obtained from the apoenzyme from both hyperthermophiles. The structure of the M. kandleri apoenzyme was determined by the multiple anomalous dispersion (MAD) method based on selenomethionine-labelled protein at medium resolution [31]. Subsequently, the structure of the M. jannaschii apoenzyme was solved at 1.75 A˚ using the molecular replacement method for phase determination [31]. A year later the structure of the holoenzyme reconstituted from the heterologuously produced apoenzyme from M. jannaschii and FeGP cofactor was solved to 1.75 A˚ resolution [11].

4.3. Protein Fold [Fe]-hydrogenase is a homodimer with dimensions of 90 A˚50 A˚40 A˚ that can be subdivided into a central globular unit and two peripheral globular units which are linearly aligned. The central unit is composed of the intertwined C-terminal segments of both subunits, forming a novel intersubunit fold. The two peripheral units consist of the N-terminal domain of each subunit and are composed of an a/b structure that belongs to the Rossmann fold family. A deep cleft is formed between both units that is built up from segments of the peripheral unit of one subunit and segments of the central unit of the other subunit (Figures 4 and 5).

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Figure 4. [Fe]-hydrogenase apoenzyme (A) from M. jannaschii and (B) from M. kandleri. The active site cleft is in a closed conformation in A and in an open conformation in B [31].

4.4. The Binding Sites for the Iron Guanylylpyridinol Cofactor and for Methenyl-H4MPT1 The Rossmann fold-like structure of the N-terminal domain contains a mononucleotide binding site for the GMP moiety of the FeGP cofactor with the binding motif GCG (Gly7, Ala8, and Gly9), which is only the first part of the characteristic dinucleotide binding motif G(X)XGXXG reflecting that the FeGP cofactor is a phosphodiester rather than a diphosphate. The other binding site for the cofactor is Cys176 which is located at the bottom of the intersubunit cleft. Cys176 is absolutely essential for enzyme activity [6] and therefore considered as iron ligand. Structural studies on the holoenzyme confirmed the previously predicted binding site. The binding site for the substrate methenyl-H4MPT1 could, so far, not be structurally characterized, but it is highly likely that it is embedded into the intersubunit cleft where sufficient space is available for the bulky molecule. In this position methenyl-H4MPT1 can be modelled in a manner that the methenyl carbocation (C14a) is near the iron site such that H2 can interact with the iron and attack C14a of methenyl-H4MPT1 from the Re-face [31] (Figure 1). Methenyl-H4MPT1 binding is presumably accompanied by a large induced-fit movement which optimizes substrate binding Met. Ions Life Sci. 2009, 6, 219–240

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Figure 5. Electron density (in blue) of the FeGP cofactor bound to [Fe]-hydrogenase of M. jannaschii. (A) Ribbon diagram of the [Fe]-hydrogenase holoenzyme; (B) Magnification of the electron density; (C) Fit of the electron density to the FeGP cofactor [11].

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and completely shields the active site from bulk solvent. Different orientations between the central and peripheral units dependent on substrate binding is reflected in the crystal structures determined [11,31]. The cleft is in a more closed state in the M. jannaschii apoenzyme structure but in an open state in the M. jannaschii holoenzyme and M. kandleri apoenzyme structures [31]. The two states can be interconverted by rotation of the peripheral units relative to the central subunit by 351 (Figure 4) [31]. Furthermore, a relatively large conformational change is indicated by the finding that [Fe]-hydrogenase crystals immediately crack upon soaking with methenyl-H4MPT1. Also it was found that in the presence of methenylH4MPT1, [Fe]-hydrogenase does not crystallize under the same conditions under which the enzyme crystallizes in the absence of methenyl-H4MPT1.

5. LIGANDS TO IRON IN THE ACTIVE SITE OF [Fe]HYDROGENASE The nature of the iron ligands and their spatial arrangement were identified by IR [29], Mo¨ssbauer [30], XAS [6], and NMR [28] spectrocopic methods and by the X-ray structure of the holoenzyme [11].

5.1. Intrinsic Ligands In the crystal structure (Figure 5) the electron density of FeGP is well shaped except for the carboxyl group of the carboxymethyl group of the pyridine which is partially disordered [11]. Accordingly, the iron is coordinated by five or six ligands forming a slightly distorted square pyramid or an octahedral arrangement. The iron is ligated by two CO ligands originally derived from IR spectroscopic data and later also seen in the crystal structure. The third iron ligand is the nitrogen and not the hydroxyl group or the carboxylate group of the pyridone which is in its pyridinol tautomeric form as deduced from the [Fe]-hydrogenase holoenzyme structure. The guanylylpyridinol ligand, especially when its hydroxyl group is negatively charged, is predicted to have back-bonding properties similar to a cyanide ligand, which is of interest since in [FeFe]-hydrogenases and [NiFe]-hydrogenases the active site iron is decorated with CO and cyanide ligands. As previously found in the latter hydrogenases the pyridinol nitrogen and the two CO molecules of [Fe]-hydrogenase are oriented perpendicular to each other. The fourth ligation site of iron is occupied by the thiolate group of Cys176 which was assumed from results of site-specific mutagenesis experiments and clearly verified in the crystal structure [11]. Met. Ions Life Sci. 2009, 6, 219–240

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The structure of the cofactor bound in the [Fe]-hydrogenase holoenzyme differs from that proposed for the protein-free cofactor (Figure 2) in that Cys176 from the apoenzyme rather than mercaptoethanol provides the sulfur ligand and in that the carboxymethyl group of the pyridinol is not involved in iron ligation. At the position of the carboxymethyl group in the protein-free cofactor an unknown ligand binds in the holoenzyme whose electron density is clearly connected with iron. The iron coordination is in good agreement to that calculated from XAS spectra which indicate that the iron in the reconstituted holoenzyme is tetra-/penta-coordinated (one sulfur, two CO, and one/two N/O) and in the protein-free cofactor it is pentacoordinated (one sulfur-, two CO, and two N/O ligands) [6]. In the crystal structure of the reconstituted holoenzyme the vacant sixth coordination site contains a spherical electron density interpreted as monoatomic solvent molecule that is, however, at a distance of 2.7 A˚ too far away to be considered as a ligand [11]. This could be the binding site for a proton or H2 which is not detectable with the methods employed (X-ray diffraction and XAS). The sixth coordination site is definitely the binding site of the competitive inhibitor CO (see below). Differences in coordination do not result in differences in the spin or oxidation state which always remains low spin Fe(II) or Fe(0) as revealed by Mo¨ssbauer spectroscopy of 57 Fe-labelled samples [30].

5.2. Carbon Monoxide and Cyanide as Extrinsic Ligands [Fe]-hydrogenase is reversibly inhibited by CO. The XAS spectrum of the CO inhibited enzyme reveals three CO, one sulfur and one/two N/O ligated to the iron [6]. The three CO bands show an almost equal intensity in the FTIR spectra (Figure 6) indicating that the extrinsic CO is bound perpendicular to the two intrinsic CO [5]. This finding attributes to the external CO the sixth ligation site of the iron in the holoenzyme. As CO is a competitive inhibitor for [Fe]-hydrogenase this site is presumably also the position of H2 binding. Unfortunately, the crystal structure of the CO inhibited enzyme was almost identical to that of the uninhibited enzyme. Apparently, the extrinsic CO, which binds only very weakly and is the first to come off upon light irradiation [29], was lost during crystal structure determination. Attempts to show the binding of extrinsic CO to the iron of the proteinfree FeGP cofactor gave ambiguous results. Whereas the Mo¨ssbauer spectrum of the cofactor changed significantly when CO was added to the cofactor solution [30], the FTIR spectrum did not [29], which can presently only be explained considering that the experimental conditions were quite different. The measurements of the FTIR spectra of the protein-free cofactor in the absence and presence of extrinsic CO clearly have to be revisited. Met. Ions Life Sci. 2009, 6, 219–240


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Figure 6. Infrared spectra of the [Fe]-hydrogenase holoenzyme from M. marburgensis in the absence (upper spectrum) and presence of 12CO (lower spectrum). Reproduced from Lyon et al. [29] with permission of the American Chemical Society, copyright (2004).

Cyanide reversibly inhibits [Fe]-hydrogenase by binding to the cofactor iron as shown by Mo¨ssbauer [30], FTIR [29], and XAS [6] data, the latter exhibiting the iron as penta-coordinated (one sulfur, two CO, one cyanide, and one N/O). Cyanide is a non-competitive inhibitor for [Fe]-hydrogenase which implicates that it does not bind to the sixth ligation site. The most attractive binding site is that of the unknown ligand which is corroborated by a significantly higher electron density in the crystal structure of the cyanide-inhibited enzyme [11]. To the contrary, cyanide appears to irreversibly react with the protein-free cofactor yielding various decomposition products. However, the irreversible reaction is relatively slow and it might be Met. Ions Life Sci. 2009, 6, 219–240

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speculated that the cyanide replaces the carboxylate group at the iron and thereby weakens the Fe-pyridinol linkage.

5.3. Inactivation of [Fe]-Hydrogenase by O 2 and Cu(I) In cell extracts of methanogenic archaea [Fe]-hydrogenase is extremely sensitive towards O2, in the presence of which the activity is rapidly lost. To the contrary, purified [Fe]-hydrogenase is stable and catalytically active in the presence of air. This property indicates that in cell extracts [Fe]-hydrogenase is inactivated by a reactive O2 species rather than O2 itself, most likely by the superoxide anion radical O 2 . And indeed it was found that purified [Fe]-hydrogenase is rapidly inactivated in the presence of O2 and xanthine when also xanthine oxidase is present which catalyzes the formation of O 2 under these conditions (Figure 7). [NiFe]- and [FeFe]hydrogenases are also rapidly inactivated by O 2. Another property in common is the sensitivity of [Fe]-hydrogenase towards copper ions. The contaminating concentrations in buffers are generally sufficient to inactivate the enzyme at low concentrations, which is the

Figure 7. Inactivation of [Fe]-hydrogenase from M. marburgensis by superoxide anion radical. Open squares: Purified enzyme in 100 mM Tris/HCl pH 8.0 at 30 1C under aerobic conditions (liquid phase equilibrated with air). Filled circles: The enzyme solution was supplemented with 0.5 mM xanthine and 15 mU xanthine oxidase to generate O 2 . Reproduced from Buurman [51]. Met. Ions Life Sci. 2009, 6, 219–240


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reason why the buffers in contact with the enzyme have to be supplemented either with EDTA or citrate in order to complex the copper ions. The group or groups in the active site of Fe-hydrogenase, with which O 2 and copper ions react, are not yet known. The most attractive candidate is the sulfur ligand. If so, it indicates that the sulfur is highly reactive, which may be a property exploited for catalysis.

6. PROPOSED CATALYTIC MECHANISMS Any mechanism has to explain why [Fe]-hydrogenase catalyzed an exchange of H2 with protons of bulk water only in the presence of methenyl-H4MPT1 and why for the methenyl-H4MPT1 dependent exchange the iron in the cofactor is required. Two principally different mechanisms are presently discussed both relying on the finding that C14a of methenyl-H4MPT1, when bound to [Fe]-hydrogenase, has carbocation character [32–34]. In the first mechanism the iron in [Fe]-hydrogenase is assumed to have the function of a Lewis base and in the second mechanism the function of a Lewis acid [13].

6.1. H2 Binds First to the Carbocationic C14a of Methenyl-H4MPT1 Carbocations can bind H2 either side-on or end-on. It is therefore assumed that the first step in the catalytic mechanism of [Fe]-hydrogenase is the binding of H2 to 1C14a of methenyl-H4MPT1 [35,36] (reaction 7). The bound H2 is positioned such that it can interact with the cofactors, iron functioning as a Lewis base. Via the interaction of H2 with both the carbocation and the Lewis base the H-H bond is polarized such that it is heterolytically cleavaged, the hydride reacting with the carbocation and the proton with the iron (reaction 8), where the proton is in exchange with protons of bulk water (reaction 9). The activation barrier for the reaction has been calculated to be reasonable only when in the transition state H2 binds end-on to the carbocation and the base is positioned relative to H2 such that it can directly accept the proton. (Actually, in the calculation the base was assumed to be the amine group of lysine or the carboxyl group of an aspartate or glutamate rather than a transition metal) [37–39]. The proposed mechanism predicts that any H2/H1 exchange catalyzed by [Fe]hydrogenase should be absolutely dependent on the presence of methenylH4MPT1, which is what was found. Met. Ions Life Sci. 2009, 6, 219–240

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C14a þ H2 Ð ½H2 -C14a þ

ð7Þ

½H2 -C14a þ þ ½Fe Ð H-C14a þ H-½Feþ

ð8Þ

H-½Feþ þ H2 O Ð ½Fe þ H3 Oþ

ð9Þ

þ

In Eqs. (7)–(9) 1C14a is used as abbreviation for methenyl-H4MPT1, H-C14a for methylene-H4MPT and [Fe] for the iron in the FeGP cofactor. Chemical precedent for such a pathway has recently been provided [40]. The proposed mechanism is consistent with the inhibition of [Fe]-hydrogenase by CO and by cyanide, whose binding to the iron site is expected to significantly affect its Lewis basicity. Competitive CO inhibition relative to H2 has been interpreted in a manner that H2 binds to an open coordination site rather than to a free electron pair although competitive inhibition only indicates that the binding of the competitors is mutually exclusive. The function of Fe in [Fe]-hydrogenase as Lewis base implies that in the catalytic cycle an Fe(II) hydride or an Fe(IV) hydride is formed ([H-Fe]1 in reactions 8 and 9), dependent on whether the iron before protonation is in a Fe(0) or Fe(II) oxidation state [41–43]. Since mononuclear Fe(IV) hydrides are, for thermodynamic reasons, not stable, this mechanism would favor an Fe(0) complex as proton acceptor. The formation of metal hydrides in the active site of enzymes is not without precedents. A Ni(III) hydride has recently been characterized as intermediate in the catalytic cycle of [NiFe]-hydrogenase [44]. A Ni(III) hydride was also shown to be formed when the Ni(I) in methyl-coenzyme M reductase interacts with coenzyme M in the presence of coenzyme B [45]. These findings indicate that transition metals in enzymes can have Lewis base character. Despite all these pros, the mechanism fails until now to explain why the base required has to be an iron and cannot simply be a proton accepting group of the protein.

6.2. H2 Binds First to the Active Site Iron Carbonyl Complex Iron carbonyl complexes with an open coordination site form side-on (Z2-H2)Fe complexes, in which the pKa of H2 is lowered from 35 to below 15 [46–48] and in which H2 can be in proton exchange with bulk water [49,50]. Such a Lewis acid function is proposed for the iron in [Fe]-hydrogenase in the second mechanism [13]. In the first step H2 binds side-on to the iron (reaction 10) but binding is only weak with the consequence that the pK is Met. Ions Life Sci. 2009, 6, 219–240


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lowered only somewhat and that therefore the proton exchange rates are too low to be detected by the methods employed. This has to be proposed since in the absence of methenyl-H4MPT1 [Fe]-hydrogenase does not catalyze such an exchange. After H2, methenyl-H4MPT1 binds such to the enzyme that its carbocationic C14a is juxtapositioned to the iron allowing the interaction of H2 with both the iron and 1C14a (reaction 11). The interaction would lead to a polarization of H2 to an extent in which a bridging hydride and a bridging proton are formed between the two nucleophiles. The cationic (m-H2) complex thus formed is predicted to be a relatively strong acid and therefore to be in exchange with bulk water (reaction 12). ½Fe þ H2 Ð ðZ2 -H2 Þ½Fe 14a

ðZ2 -H2 Þ½Fe þ þ C

Ð ðm-H2 Þ complexþ

ðm-H2 Þ complexþ þ H2 O Ð ðm-HÞ complex þ H3 Oþ

ð10Þ ð11Þ ð12Þ

7. CONCLUDING REMARKS A crystal structure of [Fe]-hydrogenase with its substrate methenylH4MPT1 bound is not yet available. It is therefore not known, whether or not the Fe of the FeGP cofactor and the C14a of methenyl-H4MPT1 are positioned in the active site of the enzyme such that H2 can simultaneously interact with both the iron and the carbocationic C14a. For density function theory calculations all coordinates will have to be known. Model complexes to be constructed on the basis of the iron center of [Fe]-hydrogenase should give further insight into the essential but not yet understood function of the low spin iron with its unique ligands in H2 activation.

ACKNOWLEDGMENTS This work was supported by the Max Planck Society and by the Fonds der Chemischen Industrie.

ABBREVIATIONS C14a EDTA F420 1

methenyl group of methenyl-H4MPT1 ethylenediamine-N,N,N 0 ,N 0 -tetracetate coenzyme F420 Met. Ions Life Sci. 2009, 6, 219–240

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FeGP cofactor Frh FTIR GMP GP H4MPT H-C14a Hmd HPLC MAD MALDI TOF MS Mtd NMR XAS


iron guanylylpyridol cofactor of [Fe]-hydrogenase F420-reducing [NiFe]-hydrogenase Fourier transform infrared guanosine 5 0 -monophosphate guanylylpyridol tetrahydromethanopterin methylene group of methylene-H4MPT Hydrogen forming methylenetetrahydromethanopterin dehydrogenase, [Fe]-hydrogenase high performance liquid chromatography multiple anomalous dispersion matrix assisted laser desorption/ionisation time-offlight mass spectrometry methylenetetrahydromethanopterin F420-dependent dehydrogenase nuclear magnetic resonance X-ray absorption spectroscopy

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8 The Dual Role of Heme as Cofactor and Substrate in the Biosynthesis of Carbon Monoxide Mario Rivera and Juan C. Rodrı´guez Ralph N. Adams Institute for Bioanalytical Chemistry, Department of Chemistry, University of Kansas, Multidisciplinary Research Building, 2030 Becker Dr., Lawrence, KS 66047, USA <[email protected]> <[email protected]>

ABSTRACT 1. INTRODUCTION 1.1. Carbon Monoxide: Properties and Environmental Sources 1.2. The Two Faces of Carbon Monoxide: Toxic and Cytoprotective Effects 1.3. Heme Oxygenase Is a Ubiquitous Enzyme 2. THE BIOSYNTHESIS OF CARBON MONOXIDE 2.1. Heme Breakdown and Carbon Monoxide Release. Overview of the Heme Oxygenase Catalytic Cycle 2.2. The Structure of Heme Oxygenase 2.3. Formation of Hydroperoxide at the Catalytic Center of Heme Oxygenase 2.4. A Conserved Network of Hydrogen Bonded Waters Facilitates Formation of the Ferric Hydroperoxide Intermediate 2.5. Oxidation of Meso-Hydroxyheme to Verdoheme and the Release of Carbon Monoxide Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00241

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3. HEME OXYGENASE FAVORS HEME HYDROXYLATION OVER FERRYL FORMATION. THE NATURE OF THE FERRIC HYDROPEROXIDE COMPLEX IN HEME OXYGENASE 3.1. Studies with Model Heme Complexes 3.2. Studies with the Hydroxide Complex of pa-Heme Oxygenase (pa-HO-OH) 3.3. Studies with the Azide Complex of pa-HO (pa-HO-N3) 3.4. A Hydrogen Bond from Gly125 N-H to Coordinated Azide-N May Promote the Unusual Electronic Structure of the pa-HO-N3 3.5. Implications to the Mechanism of Heme Oxidation by Heme Oxygenase 4. HEME OXYGENASE DYNAMICS AND HEME BREAKDOWN. THE DISTAL LIGAND HAS A PROFOUND EFFECT IN THE DYNAMIC BEHAVIOR OF HEME OXYGENASE 4.1. H/D Exchange 4.2. Microsecond-Millisecond Dynamics 4.3. A Unifying View of Protein Dynamics and Heme Oxygenase Reactivity 5. THE REGIOSELECTIVITY OF HEME HYDROXYLATION 5.1. Hydroxylation of the a-Meso Carbon Leads to Its Release as CO with Subsequent Formation of a-Biliverdin 5.2. pa-Heme Oxygenase Exhibits Unique d-Regioselectivity 5.3. Polypeptide-Heme Interactions Control the Regioselectivity of Heme Oxidation 5.4. The 1H NMR Spectra of Cyanide-Inhibited Heme Oxygenase as a Diagnostic Tool of Heme Oxidation Regioselectivity 6. CONCLUSION AND OUTLOOK ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

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ABSTRACT: Carbon monoxide (CO) is a ubiquitous molecule in the atmosphere. The metabolism of mammalian, plastidic, and bacterial cells also produces CO as a byproduct of the catalytic cycle of heme degradation carried out by the enzyme heme oxygenase (HO). The biological role of CO spans the range from toxic to cytoprotective, depending on concentration. CO generated by the catalytic activity of HO is now known to function in several important physiological processes, including vasodilation, apoptosis, inflammation, and possibly neurotransmission. Consequently, understanding the details of the reaction that leads to the formation of this important gaseous molecule from heme has become an important aspect in the study of the chemistry and

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biochemistry of HO, which utilizes heme in the dual capacity of substrate and cofactor. In this chapter, a summary, and when appropriate, discussion of the current understanding of the structural, dynamical, and reactive properties that allow HO to breakdown heme into iron, biliverdin, and CO is presented. KEYWORDS: carbon monoxide enzyme dynamics ferric hydroperoxide heme degradation heme oxygenase NMR spectroscopy of heme oxygen activation protein dynamics

1. INTRODUCTION 1.1. Carbon Monoxide: Properties and Environmental Sources Carbon monoxide is a colorless and odorless gas under atmospheric temperature and pressure, with a melting point of 205 1C, a boiling point of 191.5 1C and a density of 1.250 g/L at 0 1C [1]. CO is a ubiquitous atmospheric component because it is generated from the incomplete burning of biomass (wild fires), it is released from the earth’s mantle, where it is dissolved in molten rock, by volcanic eruptions, and is produced by oxidation of methane emitted from wetlands and ruminants, and from the oxidation of higher hydrocarbons emitted by vegetation [2]. CO in the atmosphere is also a byproduct of human activity because the molecule is also produced from the incomplete combustion of fuels in households and in the now omnipresent internal combustion engine. As of 2001, the combined emission of CO is estimated to be 2780 teragrams per year; approximately 50% of the total is thought to be contributed by human endeavors and the remainder by the natural sources mentioned above [3,4]. In addition, cellular metabolism in mammals, plants and bacteria produce CO, mainly from the degradation of heme by the enzyme heme oxygenase [5–8].

1.2. The Two Faces of Carbon Monoxide: Toxic and Cytoprotective Effects Similar to several other molecules, the effect of CO on living organisms can range from toxic to essential, depending on dose. The toxicity and lethality of CO at elevated concentrations has been recognized as early as the mid 1800s, when it was identified as a poisoning agent in coal gas. Since then a very large number of confirmed cases involving CO poisoning have been documented [9]. The largest number of unintentional fatalities is associated with automobile exhaust, followed by the burning of wood and coal, and the use of kerosene stoves in improperly ventilated spaces [2]. Clinical signs of Met. Ions Life Sci. 2009, 6, 241–293

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acute CO toxicity are not specific but include dizziness, headache, weakness, confusion, abdominal pain, and muscle cramping. Several of these symptoms are consistent with hypoxia, which is a consequence of the fact that carbon monoxide competes effectively with molecular oxygen (O2) for binding to hemoglobin (Hb), thus forming carboxyhemoglobin (Hb-CO). The binding affinity of hemoglobin for CO is approximately 200 times larger than the affinity for O2 [10], which allows a gradual outcompeting of O2 for one or more of the four heme molecules in the tetrameric hemoglobin. Partial saturation of hemoglobin with CO has important implications on its affinity for O2, a fact that is manifested in a left shift of the oxygen-hemoglobin dissociation curve, which is left shifted and is accompanied by a change in shape from sigmoidal to hyperbolic. These changes in the dissociation curve indicate that partial saturation of Hb with CO impair the release of O2 at the tissue level, thus ensuing hypoxia [11]. It is important to note, however, that CO binding to hemoglobin does not account for all the pathophysiologic consequences of CO-induced toxicity, and that carbomonoxy-hemoglobin levels can be valuable in confirming CO exposure, although these levels alone cannot be used to assess the severity of the exposure or to guide a treatment plan [12]. Interestingly, animals transfused with blood containing highly saturated Hb-CO but minimal free CO did not show clinical symptoms, an observation that has led to the suggestion that a small fraction of CO dissolved in plasma may play an important role in mediating CO toxicity [9,13]. In this context, it is interesting that approximately 15% of the total CO absorbed is bound to extravascular heme proteins and enzymes such as myoglobin, cytochrome P450, cytochrome c oxidase, soluble guanylate cyclase, inducible nitric oxide synthase, and the heme-heme oxygenase complex. Taken together, these observations led to the idea that CO toxicity is a combination of tissue hypoxia and direct CO-mediated cellular damage [9]. In fact, although a complete understanding is still lacking, there are some clues indicating that cellular damage may be mediated by interactions between these heme proteins and CO. For instance, it has been shown that the activity of cytochrome c oxidase (complex IV of the mitochondrial respiratory chain) is partially inhibited upon exposure to CO, likely due to the binding of CO to the heme a3 [14] and binding of CO to the distal site of cytochromes P450 inhibits the activity of these widespread and important enzymes [15]. In comparison to its toxic and lethal side, the beneficial properties of CO have been recognized very recently. In fact, this has become an area of intense investigation, where new observations and plausible mechanisms of action span the range from fundamental studies to possible clinical applications. The interested reader is referred to exhaustive reviews addressing the role of CO in biology and medicine [16–19]. Carbon Met. Ions Life Sci. 2009, 6, 241–293


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monoxide in mammals is generated by the enzymatic activity of at least two isoforms of the enzyme heme oxygenase (HO), the inducible HO-1 and the constitutively expressed HO-2; the existence of a third isoform, HO-3 remains controversial [19]. Although, the endogenous synthesis of carbon monoxide had been suspected for a long time on the basis that patients with hemolytic anemia exhaled air with relatively high concentrations of CO, the biochemical basis for CO biosynthesis was not set in place until 1968, when Tenhunen, Marver, and Schmid established the existence of a previously undescribed microsomal enzyme capable of degrading heme to biliverdin [20]. In the subsequent year these investigators established that this enzyme, now known as heme oxygenase, uses NADPH and O2 to oxidize heme to biliverdin, with the concomitant release of CO and iron [5]. In vitro and in vivo studies have shown that CO generated endogenously by the activity of HO plays a role in a number of physiological processes, including vasodilation, inhibition of smooth muscle cell proliferation, inhibition of apoptosis, and possibly in neurotransmission. These cytoprotective properties of CO have been reviewed exhaustively [16–19]. Endogenously generated CO also appears to have an antiinflammatory effect, although its role independent of the participation of other byproducts of heme degradation (biliverdin and up-regulated ferritin), is not yet clear. The possibility that CO generated by induction of HO-1 may exert antiinflammatory effect has been mirrored in the antiinflammatory response induced by exogenous CO, which was released from administered methylene chloride upon enzymatic degradation [21]. These findings, which include protection of grafts in animal studies of organ transplantation [21,22], stimulated some investigators to explore the idea of synthesizing compounds capable of delivering CO to tissues, with the expectation of using them as potential antiinflammatory drugs [23–25].

1.3. Heme Oxygenase Is a Ubiquitous Enzyme The degradation of heme is catalyzed by the enzyme heme oxygenase [5], which carries out a complicated set of reactions that result in opening of the heme macrocycle, release of the heme iron, and production of biliverdin and carbon monoxide (Figure 1). HO-1 has been the most actively investigated isoform because it is induced by numerous stimuli such as heme, metals, hormones, and oxidizing agents [18]. HO-2 is a constitutively synthesized enzyme present in highest concentration in the testes and brain [18]. Not long ago, the catalytic activity of HO was regarded only in the context of the maintenance of cellular heme homeostasis as a catabolic enzyme, and the products of HO activity were thought of as toxic waste material. This view changed drastically once it became apparent that the products of heme Met. Ions Life Sci. 2009, 6, 241–293

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Figure 1. Schematic representation of the heme oxidation path, leading to the formation of CO, iron, and biliverdin.

degradation play important biological functions. In addition to the now recognized importance of CO in biology and medicine (see above), the other two products of HO activity are known to have important physiological roles. Thus, heme breakdown by HO is crucial for the recycling of iron because only 1-3% of the daily iron requirement is obtained from dietary intakes [26] and biliverdin and bilirubin are powerful antioxidants [27–29]. Heme oxygenase has also been identified in some pathogenic bacteria. In this context, it is interesting that a primary obstacle for successful bacterial colonization of a mammalian host (infection) is the lack of available iron because the concentration of free iron in mammals is maintained at a very low level, B109 M [30]. Some bacterial pathogens are capable of utilizing heme as a sole source of iron, suggesting that bacterial HOs are integral part of a pathway to mine iron from host heme. Nevertheless, a relatively small number of bacterial heme-degrading enzymes have been identified so far [6,31–35]. The biochemical and biophysical characterization of these enzymes has provided an additional avenue to explore the mechanism of heme catabolism at the molecular level. Heme oxygenase also plays an important role in plants where it facilitates the synthesis of phycobilins and open-chain tetrapyrroles, which are produced from the final product of heme degradation, biliverdin [7,8]. Heme oxygenase enzymes have been found to be essential in the cyanobacterium Synechocystis sp. PCC6803, the red algae Cyanidium caldarium [36], and in the higher plant Arabidopsis thaliana [37], thus HO seems to be present ubiquitously in the plant kingdom. A recent study reported the bacterial expression and spectroscopic characterization of heme oxygenase from Synechocystis sp. PCC6803 [38] (Syn HO-1) and concluded that the heme binding site of this enzyme is likely to be more similar to that of the bacterial Met. Ions Life Sci. 2009, 6, 241–293


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cd-HO than to the mammalian enzymes. Future detailed characterization of plant HOs is also likely to prove fruitful in the quest to understand the mechanism of heme degradation. There are several excellent reviews summarizing the rich chemistry and biochemistry [39–43] and the biology [18,28,44,45] of heme oxygenase enzymes, thus this chapter is in no way a comprehensive recapitulation of the immense field of heme oxygenase research. Rather, it will focus on the structural, dynamic and reactive properties inherent in heme oxygenase that allow this enzyme to channel dioxygen activation toward heme oxidation, which leads to the biosynthesis of CO, with concomitant production of iron and the antioxidant biliverdin.

2. THE BIOSYNTHESIS OF CARBON MONOXIDE 2.1. Heme Breakdown and Carbon Monoxide Release. Overview of the Heme Oxygenase Catalytic Cycle Heme oxygenase is unusual in that it utilizes heme in a dual role of substrate and cofactor. It is now clear that the catalytic cycle of HO (Figure 1) parallels that of cytochrome P450 in that the ferric enzyme is reduced to its ferrous state, with subsequent formation of an oxyferrous complex (FeII-O2) that accepts a second electron and is thereby transformed into an activated ferric hydroperoxy (FeIII-OOH) oxidizing species [46]. Thereafter the mechanisms of heme hydroxylation and monooxygenation diverge, as shown schematically in Figure 2. Whereas the FeIII-OOH oxidizing species in monooxygenases decays to an oxoferryl species (FeV¼O), the FeIII-OOH intermediate in HO reacts with the heme macrocycle to give a-meso hydroxyheme. The latter undergoes a subsequent O2-dependent elimination of the hydroxylated a-meso carbon as CO with the concomitant formation of verdoheme (see Figure 1). Verdoheme is subsequently oxidized to FeIII-biliverdin in a reaction that requires both O2 and reducing equivalents [46,47]. In mammals, FeIII-biliverdin must be reduced to FeII-biliverdin previous to the sequential release of FeII and biliverdin [48]. In total, 7 electrons and 3 molecules of O2 are needed to oxidize heme to biliverdin. Cytochrome P450 reductase (CPR) supports the catalytic activity of HO-1 by donating all 7 electrons to mammalian HO-1 enzymes [42,48]. Electron transfer from CPR to HO is preceded by the formation of a ternary complex involving CPR, NADPH and HO; the binding interface between HO-1 and CPR has been partially delineated using results obtained from fluorescence resonance energy transfer (FRET) [49] and surface plasmon resonance (SPR) studies [50]. In the case of bacterial HOs, it has recently been Met. Ions Life Sci. 2009, 6, 241–293

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Figure 2. The obligatory FeIII-OOH intermediate is common to monooxygenase (cyt P450) and peroxidase enzymes, in which it decays to a formally FeV oxo oxidizing species. The FeIII-OOH intermediate is also obligatory in heme oxygenase where it reacts with the heme to form meso-hydroxyheme.

established that the electron donor to HO from P. aeruginosa (pa-HO) is an NADPH-dependent ferredoxin reductase, pa-FPR [51]. The genes coding for pa-HO and pa-FPR are upregulated when P. aeruginosa is challenged with low iron concentrations [52], an observation that underscores the synergistic function of these enzymes in the release of iron from heme [51].

2.2. The Structure of Heme Oxygenase The crystal structure of human HO-1 (h-HO-1), first reported in 1999 [53], catalyzed an intense spark of activity that allowed relatively fast progress toward the fundamental understanding of HO chemistry and biophysics. This seminal report was rapidly followed by the elucidation of the X-ray crystal structure of rat heme oxygenase-1 (r-HO-1), which is 84% identical to h-HO-1 [54]. More recently, the crystal structures of heme oxygenase enzymes from pathogenic bacteria such as Neisseria meningitidis (nm-HO) [55], Corynebacterium diphtheriae (cd-HO) [56] and Pseudomonas aeruginosa (pa-HO) [57] have been solved. Despite the low sequence identity of nm-HO (22%) [32,58], cd-HO (33%) [31], and pa-HO (19%) [33] relative to HO-1 the bacterial HO enzymes share the fold characteristic of the mammalian enzymes. The HO fold is highly a-helical (8 helices) and harbors the heme between helices A and F, as shown in Figure 3. This fold is also shared Met. Ions Life Sci. 2009, 6, 241–293


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by the constitutively expressed mammalian enzyme HO-2 [59,60]. Human HO-2 is one of several HO enzymes whose structure has been solved in its substrate-bound and apo forms. In all cases, the overall fold of the holo-enzyme is maintained in the absence of heme [59,61]. A common feature among all known holo-HO structures is the presence of several water molecules encased between helix C, E, and F at the core of the protein. These structural water molecules, which are detected in X-ray diffraction maps, can also be inferred from their NMR spectroscopic characteristics in solution [62], and constitute an integral part of a hydrogen bonding network that is known to play essential catalytic roles in HO. Site-directed mutagenesis and spectroscopic studies conducted before the structure of HO-1 was available suggested that the heme environment in the heme-HO complex is similar to that of myoglobin in that its heme is coordinated by a proximal, non-ionized His residue (yellow in Figure 3) and by a distal H2O or OH ligand [63–65]. The advent of the crystal structure of HO-1 [53] revealed the absence of polar residues in the distal binding site, and thus the absence of a distal residue. In fact, as can be seen in the view of Figure 4, the backbone of a section of helix F is in close contact with the heme, a structural motif that is unusual in heme proteins. Consequently, the

Figure 3. X-ray crystal structure of h-HO-1 (PDB ID 1N45) showing the a-helical fold characteristic of HO enzymes. The heme (red) is sandwiched between helices A and F and is coordinated by a proximal histidine (yellow). Met. Ions Life Sci. 2009, 6, 241–293

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Figure 4. Blow-out of helices A and F in pa-HO to illustrate that the distal helix (F) above the heme does not contain polar side chains. It is also apparent that the backbone of helix F makes contact with the face of the heme. This is unusual in heme proteins but characteristic of HO enzymes of known structure.

only polar groups near the heme-iron that can stabilize the distal H2O or O2 ligands are the carbonyl oxygen of Gly139 and the amine nitrogen of Gly143 [53]. The crystal structure of rat HO-1 [54] also showed that Gly139 and Gly143 are near the heme iron and likely contribute to stabilizing the distal H2O or O2 ligands. The authors of this study also suggested that the N-H group of Gly143 may form a hydrogen bond to the coordinated oxygen atom of the hydroperoxide molecule in the FeIII-OOH oxidizing species, thus assisting in the catalysis of heme hydroxylation [54]. We will return to this issue latter in this chapter. The fold of bacterial HOs is almost identical to that of their mammalian counterparts. The proximal side of the heme pocket in the bacterial enzymes also harbors a non-ionized His ligand [55]. Interestingly, but perhaps not surprisingly given the low sequence similarity, the amino acid identity among residues comprising the distal helices of mammalian and bacterial HO enzymes is low. This is manifested in several structural differences in the distal helix, where there is no conservation of acidic and basic side chains, which would appear to eliminate the possibility of a conserved role in sustaining catalysis for these types of residues [55]. Nevertheless, it is noteworthy that Gly116 and Gly120 in nm-HO, Gly135 and Gly139 in cd-HO, and Gly121 and Gly125 in pa-HO take the place of Gly139 and Gly143 in the distal pocket of HO-1. It has therefore been suggested that the distal helix kink and flexibility imparted by these two Gly residues is an important Met. Ions Life Sci. 2009, 6, 241–293


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component of HO function [55]. Hence, although the distal helix of HO enzymes can tolerate significant differences in amino acid composition and structure, the presence of Gly139 and Gly143 (h-HO-1), or their equivalent in distinct heme oxygenase enzymes, is necessary to support heme hydroxylation. Consistent with this idea, when Gly139 or Gly143 in HO-1 is replaced, the FeIII-OOH intermediate in the mutant enzymes does not attack the heme to form meso-hydroxyheme but rather decays into a ferryl species [66]. Recent reports suggest that the product of the ChuS gene in pathogenic E. coli O157:H7 is a heme oxygenase possessing a fold distinct from that displayed by all HOs of known structure [67,68]. In contrast to the classical a-helical fold of HO, the structure of ChuS consists of a central core made up of two b-sheets, each consisting of nine antiparallel b-strands flanked by a pair of parallel b-helices. It is interesting that although the structure of ChuS consists of two identical tandem repeats, the structure of the heme complex shows that heme binds only at the C-terminal motif. Clearly, additional work is needed to establish whether the ChuS fold is mirrored in HO enzymes from other bacterial strains.

2.3. Formation of Hydroperoxide at the Catalytic Center of Heme Oxygenase The activation of O2 by HO starts with the one-electron reduction of the heme-HO complex, followed by the binding of dioxygen to form an oxyferrous complex (FeII-O2) [46], as shown schematically in Figure 1. The efficient utilization of O2 depends in part on the formation of a stable FeII-O2 complex because unstable complexes are not only expected to favor the dissociation of O2 but would also be susceptible to autoxidation, thus lowering the efficiency of O2 utilization. Indeed, it has been shown that the affinity of ferrous HO-1 and HO-2 for O2 is 30 mM1 and 80 mM1, respectively, which is 30–90-fold greater than that exhibited by mammalian myoglobins [69]. The enhanced stability of the oxyferrous complex in HO is manifested in relatively high O2 association rate constants (similar to those exhibited by myoglobin) and in significantly slower O2 dissociation rates, which are notably slower than those measured for myoglobin. The relatively high affinity for O2 also imparts HO with a relative immunity toward inhibition by the CO formed during heme oxidation. This point will be addressed in more detail in Section 2.5. Early work with heme oxygenase established that the oxidation of heme to biliverdin requires that the FeII-O2 complex accepts a second reducing equivalent [46]. This information was interpreted to suggest that the Met. Ions Life Sci. 2009, 6, 241–293

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reduction of the coordinated O2 molecule in the oxyferrous complex of HO leads to the formation of a coordinated peroxide, which acts as the heme hydroxylating species [70]. The same investigators demonstrated that the addition of limited amounts of H2O2 to a solution of rat HO-1 results in the oxidation of the heme to verdoheme. Interestingly, substituting H2O2 by alkyl hydroperoxides did not result in heme hydroxylation or the formation of verdoheme. Instead, it was observed that acyl- and alkylhydroperoxides promote the formation of a ferryl intermediate [70]. The results obtained upon reacting H2O2 with HO stand in stark contrast with those obtained upon reacting H2O2 or alkylhydroperoxides with ferric porphyrinates [71] or with heme proteins such as peroxidases [72] and globins [73], which produce a ferryl complex, regardless of the nature of the oxidant i.e., H2O2 or ROOH. It was therefore concluded that the nature of the species that oxidizes the HO-bound heme to a-meso-hydroxyheme is a ferric hydroperoxide (FeIII-OOH) [70]. The oxidation of heme to verdoheme upon addition of limited amounts of H2O2 to a solution of HO-1 was subsequently observed with HO-2 [74] and with the bacterial heme oxygenases cd-HO [75] and pa-HO [76,77], thus strongly supporting the notion that hydroxylation of the heme-HO complex is fully supported by a ferric hydroperoxide (FeIII-OOH) intermediate. Spectroscopic evidence supporting the need for a one-electron reduction of the oxyferrous complex to form the FeIII-OOH, which in turn hydroxylates the heme in HO, was obtained from cryogenic ENDOR (electron nuclear double resonance) spectroscopic studies. These experiments showed that upon radiolytic reduction of a frozen solution (77 K) of the FeII-O2 complex of HO, a low-spin ferriheme complex (FeIII-OO) is formed, which rapidly accepts a proton to form the FeIII-OOH species [78]. The latter exhibits a rhombic EPR (electron paramagnetic resonance) spectrum characteristic of low-spin ferrihemes and is stable at 77 K. Upon increasing the temperature to 218 K for a short period of time and cooling back to 77 K the EPR spectrum of the FeIII-OOH intermediate is replaced by the spectrum of the a-meso-hydroxyheme complex of HO [78]. These findings confirmed the idea that a ferric hydroperoxide intermediate is a precursor of a-hydroxyheme and reinforce the notion that the oxidation of heme by a ferric hydroperoxide species is a novel reaction performed by heme proteins [39,70]. It is interesting to appreciate that theoretical studies have led to the suggestion that compound I, [FeIV¼O], is formed by O-O bond heterolysis of the FeIII-OOH intermediate, followed by attack of the released H2O to form meso-hydroxyheme [79]. Experimental studies aimed at testing this hypothesis, however, demonstrate that compound I in HO-1 does not hydroxylate its heme meso carbon, thus apparently ruling out the involvement of compound I in heme catabolism [80]. Met. Ions Life Sci. 2009, 6, 241–293


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2.4. A Conserved Network of Hydrogen Bonded Waters Facilitates Formation of the Ferric Hydroperoxide Intermediate Heme oxygenase and monooxygenase enzymes share a similar mechanism of O2 activation which leads to the formation of a ferric hydroperoxide intermediate. However, once formed, the FeIII-OOH intermediate can follow two divergent paths (schematically illustrated in Figure 2): (a) the formation of an oxoferryl species, thus avoiding the hydroxylation of heme, or (b) the efficient reaction with the porphyrin macrocycle to produce mesohydroxyheme. The existence of these two competing pathways implies that heme proteins whose primary function is that of heme catabolism favor (accelerate) the reaction between the ferric hydroperoxide intermediate and the heme macrocycle, whereas monooxygenase enzymes accelerate the decay of the FeIII-OOH complex into a ferryl species. The remainder of this section will be devoted to discuss the current state of understanding regarding the structure-function relationships that in HO result in heme hydroxylation rather than the formation of an oxoferryl. As has been pointed out earlier, the distal pocket structure in HO is significantly different from the equivalent structures in monooxygenase and peroxidase enzymes. An important signature of the distal pocket in HO is the absence of a polar side chain capable of stabilizing a coordinated H2O or O2 ligand. Stabilization of these ligands is likely accomplished by the presence of Gly143 in HO-1, or equivalent Gly in other HO enzymes, which is thought to donate a hydrogen bond to the coordinated sixth ligand [53,81]. The crystal structure of HO-1 showed that the distal helix also exhibits several side chains that although not positioned to interact directly with the distal ligand, can interact with the latter via bridging hydrogen bonding water molecules [53–54,82]. When one of these residues (Asp140) was mutated to Ala, His or Phe, the mutants exhibited less than 3% of the biliverdin forming activity characteristic of wild-type HO-1 [82]. Moreover, these mutants were found to exhibit peroxidase activity when incubated with H2O2 and guaiacol as substrates. Similar observations where made with Asp140 mutants of rat HO-1 [83]. The tantalizing observations made with Asp140 mutants of h-HO-1 and r-HO-1 led to additional investigations by ENDOR spectroscopy [78,84,85]: Radiolytic cryoreduction (77 K) of the FeII-O2 results in the formation of a FeIII-OOH species, as is the case when the FeII-O2 complex of cytochrome P450cam is radiolytically reduced at cryogenic temperatures [86]. In contrast, radiolytic cryoreduction of the FeII-O2 complex of myoglobin (not an oxygen activating heme protein) leads to the formation of a ferric peroxo (FeIII-OO) complex, which is converted to a FeIII-OOH species only when the sample is annealed to temperatures above 180 K [86,87]. These Met. Ions Life Sci. 2009, 6, 241–293

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Figure 5. Schematic representation of hydrogen bonding networks in (a) cytochrome P450cam and (b) h-HO-1.

differences in the readiness with which the FeIII-OO complex accepts a proton have been taken [84] to support the idea that O2-activating enzymes harbor a network of hydrogen bonds that is utilized to deliver a proton to the nascent FeIII-OO complex [88,89]. Along this vein, it is interesting to consider that the FeIII-OOH complex of HO does not hydroxylate the heme at 77 K. Instead, exposing this intermediate to progressively longer periods of time at 200 K, followed by cooling back to 77 K, results in slight changes in the EPR spectrum that have been attributed to rearrangements of the hydrogen bonding network in the distal pocket that lead to a reactive FeIII-OOH intermediate, denoted as R [84]. The 1H ENDOR spectrum of R revealed two signals, one originating from the proton in the FeIII-OOH moiety, denoted as H1 (see Figure 5b) and a second signal, also attributed to an exchangeable hydrogen, which has been denoted as H2. At 200 K the EPR spectrum of R and the signals of H1 and H2 in the ENDOR spectrum rapidly loose intensity, with the concomitant increase in intensity of a signal that corresponds to a-meso-hydroxyheme. In contrast, the EPR spectrum obtained upon radiolytic cryoreduction of the FeII-O2 complex of the Asp140 mutants of r-HO-1 is consistent with the formation of a FeIII-OO complex. Protonation of the latter to form the FeIII-OOH intermediate occurs only after the sample is annealed to temperatures above 180 K. In addition, and distinct from the observations made with the wild-type Met. Ions Life Sci. 2009, 6, 241–293


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enzyme, the FeIII-OOH intermediate in the Asp140 mutants is not converted to meso-hydroxyheme when annealed above 200 K; rather, it forms an EPR-silent species proposed to be the oxoferryl species [84], whose formation was inferred from electronic absorption spectroscopic studies of similar Asp140 mutants [82]. These observations clearly implicate Asp140 as an important component of the network of hydrogen bonds in the distal pocket of human and rat HO-1 that relays a hydrogen ion to the nascent FeIII-OO moiety. It is noteworthy that X-ray crystallographic studies conducted with the NO and CO complexes of nm-HO revealed that Asn118 takes the place of the catalytically essential Asp140 in mammalian HO-1 [90]. It was also found that the side chain of Arg77 moves upon ligand binding and is likely to participate in the network of hydrogen bonds that deliver a proton to the nascent FeIII-OO. Hence, it was concluded that the precise identity of the side chains involved in the hydrogen-bonding network of the distal pocket appears not to be important. On the other hand, conservation of a strong network of hydrogen bonds in the distal pocket of HO appears to be an important structural motif shared by all known HOs [90]. In this hydrogen bonding network Asp140 forms a hydrogen bond to a water molecule, which in turn forms a hydrogen bond with the terminal oxygen of the nascent FeIII-OO complex [83], as illustrated schematically in Figure 5b. The acidity of the water molecule donating the hydrogen bond (H1 in Figure 5b) to the nascent ferric hydroperoxo complex is thought to be diminished by virtue of being simultaneously hydrogen-bound to Asp140. Accordingly, the coordinated peroxide would not be fully protonated and would exhibit a decreased tendency to cleave its O-O bond, thereby allowing sufficient time for the heme hydroxylating reaction to take place [83]. In comparison, the terminal oxygen in the FeIII-OO intermediate of cytochrome P450cam is hydrogen-bonded with the side chain of Thr252 and a water molecule that interacts with the amido group of Asp251 [91], as illustrated schematically in Figure 5a. In this set of hydrogen bonds the interaction between the N-H group of Asp251 and the water molecule hydrogen-bonded to the terminal oxygen in the FeIII-OO moiety render this water molecule relatively acidic. Since theoretical investigations suggest that protonation of the FeIII-OO species results in significant weakening of the O-O bond [92,93], the hydrogen bonds to the terminal oxygen of FeIII-OO– are expected to facilitate cleavage of the O-O bond. The crystal structures of HO-1 coordinated by azide [81] and those obtained from the D140A mutant of human HO-1 in its ferric aqua, ferrous, and ferrous-NO forms [94] have demonstrated that the H-bonding network of water molecules is similar in all structures. In the structure of the D140A mutant of human HO-1, however, a new water molecule (water-3) takes the place formerly occupied by the carboxylate side chain of Asp140, thus Met. Ions Life Sci. 2009, 6, 241–293

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lifting the structural restriction imposed on water-1. This structural freedom of water-1 in the D140A mutant allows it to donate a hydrogen bond to the terminal oxygen atom of the FeIII-OOH group. Theoretical investigations suggest that protonation of the FeIII-OO species results in significant weakening of the O-O bond [92,93]. In fact, it has been observed that the O-O stretching frequency (nO-O) in FeIII-OOH is at least 50 cm1 lower than that in similar FeIII-OO, an observation that has been interpreted to indicate that a weakened O-O bond in FeIII-OOH is primed for dissociation to convert it into an oxoferryl species [95]. Consequently, donation of a hydrogen bond from water-1 to the terminal oxygen atom in the FeIII-OOH complex of the D140A mutant is expected to accelerate its decay to a ferryl moiety, thus explaining the peroxidase activity exhibited by the D140A mutant of HO-1 [94]. As is apparent from the discussion above, the crystal structures of human HO-1 [53,96] and rat HO-1 [54] suggested the presence of a number of well ordered water molecules in the distal pocket of these enzymes. Subsequent structures of bacterial heme oxygenase enzymes obtained from N. meningitidis [55], Corynebacterium diphtheriae [56], and Pseudomonas aeruginosa [57] confirmed the presence of several structural water molecules in the distal pocket of these enzymes. In addition, NMR spectroscopic studies conducted with cyanide-inhibited human HO-1 [62,97,98] and cd-HO [99] revealed the presence of exchangeable (NH and OH) resonances exhibiting strong downfield shifts (B10–17 ppm) [97]. Large downfield NH or OH 1H chemical shifts are diagnostic of the presence of very strong hydrogen bonds [100], where the backbone/side chain NH or OH protons participate as donors. The identity of these residues was obtained from resonance assignments and that of the corresponding hydrogen bond acceptors was attained from analysis of the crystal structure. These observations led to the conclusion that there is a relatively rigid network of hydrogen bonds in the distal pocket of HO that is more extensive than what was possible to infer from the X-ray crystal structures [62,97,98]. Hence, solution state investigations reinforced the idea that a hydrogen bond network, which involves Asp140 in h-HO-1 and r-HO-1, is critical to stabilizing the water molecule that donates a hydrogen bond to the nascent FeIII-OO complex.

2.5. Oxidation of Meso-Hydroxyheme to Verdoheme and the Release of Carbon Monoxide From the above discussion it is evident that the ferric hydroperoxide species in HO hydroxylates the heme to produce meso-hydroxyheme. The latter reacts with O2 to produce verdoheme (see Figure 1), with the concomitant Met. Ions Life Sci. 2009, 6, 241–293


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elimination of the hydroxylated carbon atom as CO. It has been suggested that ferric meso-hydroxyheme is converted to ferric verdoheme upon contact with O2, without the need of an electron [101]. An alternative mechanism in which ferric meso-hydroxyheme is reduced by one electron to ferrousmeso-hydroxyheme, which subsequently reacts with O2 to form ferrous verdoheme, has also been proposed [102,103]. This apparent discrepancy may be explained by the suggestion that under physiological conditions both reaction paths are likely, with one dominating over the other depending on the relative concentrations of NADPH and cytochrome P450 reductase in the cell [41]. It is noteworthy however, that there is complete agreement on the fact that in vitro the CO complex of ferrous meso-hydroxyheme is converted to the CO complex of ferrous verdoheme when the former is reacted with O2, a chemical property that is unique to verdoheme [101–103]. This unique property of the CO complex of meso-hydroxyheme was used in studies aimed at dissecting mechanistic differences between the heme oxygenation reaction, as observed with HO, and the coupled oxidation reaction [104]. The latter is a process [105] whereby heme proteins not involved in heme catabolism degrade heme to verdoheme and biliverdin, with the accompanying release of CO [106–111]. Meso-hydroxyheme exists in at least three different resonance forms: a ferric phenolate anion, a ferric keto, and a ferrous keto p neutral radical. It is interesting that the relative contribution of these different resonance structures changes upon exposure to CO because coordination of the latter to the iron stabilizes the ferrous keto p neutral radical species [112]. In general, CO binds to ferrous heme with higher affinity than O2 as is evident from the fact that the binding affinity of free heme for CO is significantly larger than its affinity for O2, as indicated by the ratio of the corresponding equilibrium association constants KCO/KO2, which is B25,000 [113]. The structural properties in the binding pocket of globins reduce this ratio to approximately 40 (see Table 1); the relatively high KCO/KO2 ratio exhibited by globins explains the fact that exposure to CO Table 1. Equilibrium association constants for the binding of O2 and CO to protoheme-X in different environments.

Protoheme Myoglobin HO-1 pa-HO nm-HO cd-HO

KO2 (mM1)

KCO (mM–1)

KCO/KO2

Reference

0.015 0.86 28 2.6 3.3 21

440 35 34–150 7.5 6.9 150

B25,000 41 1.2–5.3 2.9 2.1 7.1

[113] [119] [69] [115] [115] [174]

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results in the formation of carbomonoxy-hemoglobin and myoglobin. It is therefore interesting that CO generated from heme degradation by HO does not inhibit the enzyme, which in principle could be inhibited by binding CO at several stages of the catalytic cycle, including ferrous heme, meso hydroxyheme, and verdoheme. In fact, the relative immunity of HO to inhibition by CO was made evident in a study demonstrating that 20% CO in air is necessary to inhibit catalytic activity, which is arrested at the verdoheme stage due to the formation of a verdoheme-CO complex. Clearly this concentration of CO is significantly larger than those likely to be encountered under normal physiologic conditions [114]. A more quantitative description of the relative immunity of HO to inhibition by CO was obtained from measurements of the equilibrium association constants for the binding of CO and O2 to ferrous HO enzymes (Table 1). These investigations revealed that on average, the structure of HO enzymes lowers the magnitude of the KCO/KO2 ratio by approximately one order of magnitude relative to the KCO/KO2 value exhibited by myoglobin. The data in Table 1 also indicate that the affinity of ferrous HO-1 for O2 is approximately 30-fold larger than that of myoglobin, whereas the affinity of bacterial HOs for O2 is, on average, B5-fold larger than that of Mb. The lower affinity of the bacterial HOs (pa-HO and nm-HO) for O2 relative to HO-1 appears to be compensated by a lower affinity for CO relative to HO-1, such that the magnitude of the KCO/KO2 ratio is similar for all the enzymes; the magnitudes of KCO and KO2 for cd-HO, on the other hand, are very similar to those exhibited by HO-1. Although some important clues regarding the structural properties that allow HO enzymes to strongly discriminate against CO have emerged, a thorough understanding has not yet been obtained. The existing clues come from details in the measurement of binding affinities, which showed that although the O2 association rate constants are similar to those exhibited by the globins, the O2 dissociation rate constants are significantly slower [69,113,115]. These observations have been placed in context using information obtained from the crystal structures of HO and from insights derived from extensive work aimed at understanding the discrimination of CO by globins [116–119]. The emerging picture suggests that the slower O2 dissociation from HO enzymes stems from the ability of these enzymes to donate several hydrogen bonds to the coordinated O2. This general view is consistent with the polar nature of the distal binding site, which also harbors a network of structural waters that contribute to the stabilization of the coordinated O2. The number of hydrogen bonds and the details of the interactions, however, appear to be different in distinct HO enzymes [69,113,115]. Recent studies demonstrated that the dynamic behavior of the HO polypeptide is largely dependent on the oxidation state and on the nature of the distal ligand coordinating the heme iron [120,121] (see Section 4). It is therefore possible Met. Ions Life Sci. 2009, 6, 241–293


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that an additional source of discrimination against CO comes from the dynamic plasticity of the heme pocket in HOs, which may be different depending on whether the distal ligand is CO or O2.

3. HEME OXYGENASE FAVORS HEME HYDROXYLATION OVER FERRYL FORMATION. THE NATURE OF THE FERRIC HYDROPEROXIDE COMPLEX IN HEME OXYGENASE 3.1. Studies with Model Heme Complexes In the EPR spectrum of the ferric hydroperoxide intermediate of HO-1 [84] the sum of the squares of the g values (Sg2) is 14.14. This compressed g anisotropy (Sg2 o16) suggested [122] the possibility of an electronic configuration, (dxz,dyz)4(dxy)1, (dxy)1 hereafter [123–126]. Ferric porphyrinates exhibiting the (dxy)1 electronic configuration are known to be highly nonplanar (ruffled) and to place a large amount of unpaired electron density at the meso carbons [123–128]. In comparison, ferric porphyrinates with the more common (dxy)2(dxz,dyz)3 electron configuration, dp hereafter, exhibit planar macrocycles and place significant spin density at the a-pyrrole carbons and negligible spin density at the meso carbons [127–129]. To understand the role exerted by a hydroperoxide axial ligand on the electronic structure of ferrihemes, meso-13C labeled FeIII-tetraphenylporphyrinates coordinated by a methoxide and an alkyl hydroperoxide ligand, [meso-13C-TPPFe(OCH3)(OOtBu)], was investigated by ENDOR and 13C NMR spectroscopy. The ENDOR spectrum unequivocally indicated that the electron configuration of [meso-13C-TPPFe (OCH3)(OOtBu)] at 8 K is low-spin dp, despite the fact that the EPR spectrum displays compressed g anisotropy (Sg2 B14) [122]. In striking contrast, results obtained with 13C NMR spectroscopy at more elevated temperature suggested a different picture: The meso 13C shift of [meso-13CTPPFe(OCH3)(OOtBu)] at 218 K occurs at 422 ppm. The relevance of this observation is that ferrihemes with the (dxy)1 electron configuration exhibit large spin density at the meso carbons [127], which induce diagnostically large (B1000 ppm) meso carbon shifts [130,131]. In comparison, ferrihemes with the dp electron configuration exhibit spin density into b-pyrrole carbons [125,127] and negligible spin density at the meso carbons, which results in meso carbon resonances near the diamagnetic position, 70 ppm [132,133]. Hence, the 422 ppm chemical shift observed for [meso-13CTPPFe(OCH3)(OOtBu)] indicates significant population of the (dxy)1 electron configuration at 218 K [122]. Temperature-dependent investigations Met. Ions Life Sci. 2009, 6, 241–293


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of these resonances led the authors to suggest that a population of molecules with a dp electron configuration and planar heme is in dynamic equilibrium with a population of molecules with a (dxy)1 electron configuration and a ruffled macrocycle [122,134], as indicated by equation (1). The equilibrium shifts to the left as the temperature decreases and at 8 K the fraction with dp planar macrocycles approaches unity, which is in agreement with the results obtained from the ENDOR experiments. In contrast, at the temperature of the 13C NMR experiments the population with planar and ruffled hemes is nearly equal and at ambient temperature the population with ruffled heme and (dxy)1 electronic structure is expected to predominate [122]. The authors thus suggested that if a similar equilibrium between planar and ruffled hemes takes place in the heme pocket of HO, the relatively large electron density at the meso-carbons would facilitate attack of the terminal OH group in the FeIII-OOH complex [122]. Keq

planarðdp Þ $ ruffledðdxy Þ

ð1Þ

3.2. Studies with the Hydroxide Complex of pa-Heme Oxygenase (pa-HO-OH) The equilibrium between planar and ruffled hemes observed with the model complex described above implied that the electronic structure of the FeIII-OOH intermediate in heme oxygenase should be studied at ambient temperature [122]. In an attempt to circumvent the extremely high reactivity of this intermediate the electronic structure of the hydroxide complex of pa-HO was studied as a model of the highly reactive FeIII-OOH [135]. This approach capitalizes from methodology developed for the biosynthesis of 13 C-labeled heme [136,137] and from relatively straightforward correlations between core carbon shifts and the coordination state and electronic structure of hemes [138–140,130,132]. The 13C NMR spectra of the hydroxide complex of pa-HO (Figure 6) revealed the presence of at least three populations of molecules in dynamic exchange, each exhibiting a distinct electronic configuration [135]. The most abundant population has an S ¼ 3/2, (dxz, dyz)3(dxy)1(dz2)1 (S ¼ 3/2, (dxy)1, hereafter) electronic structure [141], which is characterized by pyrrole carbon-a (Ca) and pyrrole-b (Cb) resonances near 400 ppm and meso carbon (Cm) resonances near zero ppm [139] (blue boxes in Figure 6). The spectra also demonstrate the presence of two other populations with relatively low concentrations. One of them has an S ¼ 3/2 (dxy)2(dxz,dyz)2(dz2)1 (S ¼ 3/2, dp, hereafter) electron configuration, with Ca resonances ca. 650 ppm, Cb resonances ca. 1000 ppm and Cm resonances ca. –200 ppm (red boxes). The other low-concentration species exhibits an S ¼ 1/2, (dxy)1 electron configuration, with Cm resonances ca. Met. Ions Life Sci. 2009, 6, 241–293


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Figure 6. 13C NMR spectra (37 1C) of the hydroxide complex of pa-HO reconstituted with heme labeled at the Ca and Cm carbons (A) and Ca and Cb carbons (B). Peaks corresponding to the populations exhibiting the S ¼ 3/2, (dxy)1 spin state are in blue boxes, peaks corresponding to the population with the S ¼ 3/2, dp spin state are in red boxes, and peaks corresponding to the population with the S ¼ 1/2, (dxy)1 spin state are in black boxes. Adapted from [135].

1300 ppm and Ca resonances ca. –400 ppm (black boxes). These electronic configurations are highly unusual among heme active sites. In fact, this is the first example of a heme active site with S ¼ 3/2, (dxy)1 and S ¼ 1/2, (dxy)1 electronic configurations [135]. More important, ferric porphyrinates with these unusual electron configurations are always associated with non-planar distortions of the macrocycle [142–144,123,125,139], and also place significant unpaired electron density at the meso positions [135]. In contrast, it is noteworthy that the hydroxide complex of globins has the S ¼ 1/2, dp electron configuration typical of planar hemes [145]. Met. Ions Life Sci. 2009, 6, 241–293

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A recent 1H NMR spectroscopic study of the hydroxide complex of heme oxygenase from Neisseria meningitidis (nm-HO-OH) concluded [146] that the electronic ground state of this complex is the conventional dp orbital ground state and not the unusual S ¼ 1/2 (dxy)1 reported for the hydroxide complex of pa-HO-OH. This statement, unfortunately, only adds confusion to current understanding: The authors of the study involving pa-HO-OH did not conclude that S ¼ 1/2 (dxy)1 is the ground electronic state of this complex [135]. In fact, as can be seen in Figure 6, the 13C NMR spectra clearly indicate that the pa-HO-OH complex exists in at least 3 interconverting electronic states; the S ¼ 1/2 (dxy)1 electronic state (corresponding resonances in black boxes) is the least populated and therefore it is highly unlikely to be the electronic ground state in the pa-HO-OH complex. Thus, if the FeIII-OOH complex in HO were to behave similarly, heme hydroxylation may occur via the population with the reactive (dxy)1 electronic structure; the dynamic interconversion of populations would constantly repopulate the low abundance but reactive (dxy)1 state, therefore facilitating efficient heme oxidation. It is also important to emphasize that discovery of the unusual electronic structures in pa-HO-OH [135] was possible because this complex was studied with the aid of 13C NMR spectroscopy, in addition to the traditional approach of observing only 1H NMR resonances. Additional discussion regarding this issue is illustrative: As pointed out above, there are three 13C NMR observable populations of pa-HO-OH, each exhibiting a characteristic set of core porphyrin carbon chemical shifts [135]. The relative intensity of the peaks corresponding to the most abundant population facilitates the study of their temperature dependence (see Figure 7). Thus, it is apparent that at 37 1C (1/T B3.2 103 K1) the magnitude of Ca and Cb chemical shifts from the most abundant population is near 500 ppm and 400 ppm, respectively (D and J in Figure 7). These values are significantly larger than those corresponding to Ca and Cb in pa-HO-CN (m and K symbols), which is known to exhibit a pure low-spin dp electron configuration [141,147]. In addition, the temperature dependence of the Ca and Cb shifts from pa-HO-OH is very steep, which also stands in sharp contrast to the modest temperature dependence of the Ca and Cb shifts from pa-HO-CN. In contrast, the Cm resonances in both, pa-HO-OH and pa-HO-CN are very similar and near their corresponding diamagnetic shifts. There is no question that the information content of the 13C shifts is rich and points to the fact that the electronic structure of this most abundant population of pa-HO-OH is not the typical low-spin dp. In stark contrast, the paramagnetically resolved resonances in the 1H NMR spectrum of pa-HO-OH do not alert the experimentalist of an unusual situation. In fact, if one were to study only the 1H NMR spectrum of pa-HO-OH (Figure 8B), which is somewhat similar that of pa-HO-CN (Figure 8A), it would not be Met. Ions Life Sci. 2009, 6, 241–293


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Figure 7. Temperature dependence of core carbon chemical shifts corresponding to pa-HO-OH (open symbols) and pa-HO-CN (filled symbols). Adapted from [135].

difficult to conclude that nothing is unusual in the electronic structure of the pa-HO-OH complex. However, as discussed above, the remarkable 13C chemical shifts and their temperature dependence prompted detailed NMR spectroscopic probing of the pa-HO-OH complex [135]. These investigations revealed that the most abundant population of pa-HO-OH exists as an equilibrium mixture of two species, one exhibiting an S ¼ 1/2 dp electron configuration and planar heme and a second with a novel S ¼ 3/2, (dxy)1 spin state and nonplanar heme. As has been already mentioned, the same investigations revealed two other populations of lower concentration, one with the S ¼ 3/2, dp electronic state and the lowest concentration population with the S ¼ 1/2 (dxy)1 electron configuration.

3.3. Studies with the Azide Complex of pa-HO (pa-HO-N3) Observations made with the hydroxide complex of pa-HO suggest that the nature of the axial ligand is important in inducing the unusual heme Met. Ions Life Sci. 2009, 6, 241–293


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Figure 8.

1

H NMR spectra of pa-HO-CN (A) and pa-HO-OH (B).

electronic structures and deformations described above. For example, when CN is the distal ligand in pa-HO [147], human [148], rat [149], or bacterial [99,150] HOs, the corresponding 1H and 13C NMR spectra are always indicative of planar low-spin dp electronic hemes. In contrast, when the distal ligand is capable of accepting a hydrogen bond at the coordinating atom (i.e., OH) the resulting complex no longer exhibits the common low-spin dp electron configuration characteristic of planar ferrihemes. Thus, the azide (N 3 ) complex of pa-HO (pa-HO-N3) was studied to further probe this idea [141] because the N 3 ligand is also capable of accepting a hydrogen bond at the coordinating atom. Once again, 13C NMR spectroscopic investigations demonstrated that the pa-HO-N3 complex does not harbor a planar ferriheme with the S ¼ 1/2, dp electronic structure. Instead, the binding of azide leads to nonplanar hemes and electronic structures similar to those described above for pa-HO-OH. The magnitude and temperature dependence of the chemical shifts from the pa-HO-N3 complex [141] are strikingly similar to those exhibited by the most abundant population of pa-HO-OH (see above). Detailed analysis of these observations in the context of molecular orbitals and concomitant measurements of magnetic moment led to the conclusion that the pa-HO-N3 complex also exists at ambient temperature as a mixture of two interconverting populations, one with the dp electron configuration Met. Ions Life Sci. 2009, 6, 241–293


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and planar heme and the other with the S ¼ 3/2, (dxy)1 electron configuration and nonplanar heme [141].

3.4. A Hydrogen Bond from Gly125 N-H to Coordinated Azide-N May Promote the Unusual Electronic Structure of the pa-HO-N3 The above-described studies of the heme active site in HO led to the conclusion that the enzyme channels the activation of O2 toward heme hydroxylation aided by unusual heme electronic structures of the FeIII-OOH oxidizing species. An advantage of pa-HO-N3 as a model of the FeIII-OOH intermediate in HO is that it can be studied at neutral pH, which permits concomitant probing of the polypeptide with multinuclear and multidimensional NMR experiments. Several NMR studies of the polypeptide in HO have been carried out with the aid of homonuclear methods that rely on the larger than normal dispersion of 1H resonances affected by heme paramagnetism. These studies provided several assignments of residues in relative close proximity to the paramagnetic heme active site [151–153,62]. Nevertheless, understanding polypeptide dynamics and polypeptide-heme interactions in HO enzymes requires the availability of complete heteronuclear and sequential assignments that permit global probing of the enzyme. A study reporting the sequential assignment of backbone (N-1H, 15 N-H, Ca, Cb, and C’) resonances of the 198-residue-long, paramagnetic pa-HO in complex with azide and with cyanide paved the way to study the dynamic behavior of HOs using NMR methods [120]. The strategy used for obtaining resonance assignments from residues strongly affected by the heme iron paramagnetism made use of selective amino acid labeling and NMR experiments tailored for the observation of fast relaxing resonances [120]. Selective amino acid labeling was also used to probe the notion that the lower than typical field strength of N 3 coordinated to pa-HO likely stems from accepting a hydrogen bond at the coordinating atom. The hydrogen bond donor would be either the N-H from Gly125, either directly or mediated by a water molecule [134–135,141]. Thus, pa-HO-N3 selectively labeled with 15N Gly (15N-Gly-pa-HO) was used in these investigations [120]. Despite considerable effort, the cross peak corresponding to Gly125 was not found in the HSQC spectrum of pa-HO-N3, presumably because the proximity to the paramagnetic iron greatly increases the relaxation rate of the amide hydrogen in Gly125. On the other hand, since the gyromagnetic ratio of 15N is 10-fold smaller than that of 1H, the effect of the paramagnetic center on 15N relaxation is expected to be approximately 100-fold smaller. Met. Ions Life Sci. 2009, 6, 241–293

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Thus, at 37 1C the 15N resonance of Gly125 in pa-HO-N3 was observed at 128 ppm and 133 ppm in a one-dimensional 15N spectrum (Figure 9a) [120]. Two peaks (128 and 133 ppm) with relative intensity 70:30 are observed because pa-HO-N3 exists in solution as a mixture of two heme orientational isomers with equilibrium concentrations B70:30 [147]. Clearly, these resonances are at least 3-fold broader than other 15N resonances in pa-HO-N3, are significantly downfield shifted relative to other Gly 15N resonances, and exhibit pronounced temperature dependence (Figure 9). These unusual properties strongly suggest through-bond (Fermi contact) spin delocalization from the heme iron into the Gly125 15N atom. Through-bond spin delocalization would only be possible if the Gly125 N-H forms a hydrogen bond with the iron-bound nitrogen of azide. Additional support for this idea

Figure 9. One-dimensional 15N NMR spectrum of pa-HO-N3 labeled with 15N-Gly obtained at (a) 37 1C, (b) 32 1C, (c) 20 1C and (d) 8 1C. Adapted from [120]. Met. Ions Life Sci. 2009, 6, 241–293


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was found in the uncommonly large upfield deuterium isotope effect observed for the 15N resonance of Gly125 which also suggests unpaired spin delocalization onto the 15N atom of Gly125 via a hydrogen bond [120]. In comparison, it is important to underscore that the 15N and 1H resonances of Gly125 in pa-HO-CN, where the coordinated carbon atom cannot accept a H-bond, display ‘‘standard’’ chemical shifts, line widths, and deuterium isotope effects.

3.5. Implications to the Mechanism of Heme Oxidation by Heme Oxygenase The above-discussed observations led to the postulation [134,135,141] that coordinated OH or N 3 , by virtue of accepting a hydrogen bond to the coordinating atom, lower their s-donating ability, thereby lowering their field strength. Decreasing axial ligand field strength is typically accompanied by: (i) strengthening of the equatorial field, i.e., shorter bonds between the iron and the pyrrole nitrogens, which induce nonplanar porphyrin deformations and (ii) stabilization of the dz2 orbital which facilitates attainment of the S ¼ 3/2 electron configuration [154–156]; and/or stabilization of the dxz and dyz orbitals relative to the in-plane dxy orbital, which is conducive to the (dxy)1 electronic structure [144,157–159]. Consequently, if the field strength of the HOO– ligand is modulated similarly via a hydrogen bond to the coordinating O atom, the nonplanar deformations and unpaired spin density at the meso carbons observed in the populations with the (dxy)1 and S ¼ 3/2 spin states would facilitate homolytic cleavage of the O-O bond by efficiently trapping the OH radical at a sterically unprotected meso carbon. Thus, if a limiting resonance structure in which the electron is fully transferred from the porphyrinate ring to the metal is considered (Figure 10b), attack of OH at a meso carbon would produce c, which would rapidly rearrange its Fe-O electron configuration, lose a proton from the attacked meso carbon to complex re-aromatize the porphyrin ring, and protonate the FeIII-O2 2 to yield the ferric meso-hydroxyheme complex d [122,141]. In this context, it is interesting that a theoretical study has suggested that heme hydroxylation by HO is more likely to occur via a stepwise mechanism in which homolytic cleavage of the O-O bond is followed by trapping of the OH radical at the porphyrin [160]. Importantly, more recent calculations [161] also suggest that homolytic cleavage of the O-O bond proceeds via a species with an electronic structure resembling a porphyrin radical, Por1d-FeIIIOH---HOd. Hence, the notion derived from experimental studies with pa-HO-OH and pa-HO-N3, which suggests that the porphyrin in the FeIII-OOH intermediate has significant unpaired spin density at the meso carbons, is in good agreement with theoretical developments. Met. Ions Life Sci. 2009, 6, 241–293

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Figure 10. Schematic illustration of the proposed activation of meso heme carbons in the FeIII-OOH complex of HO. (a) The population with the (dxy)1 electronic structure can be viewed as the limiting resonance structure (b), which facilitates homolytic cleavage of the O-O bond to produce (c). The latter is expected to rapidly rearrange its Fe-O electron configuration, rearomatize the porphyrin ring and protonate the FeIII-O2 2 complex to form the meso hydroxyheme (d).

4. HEME OXYGENASE DYNAMICS AND HEME BREAKDOWN. THE DISTAL LIGAND HAS A PROFOUND EFFECT IN THE DYNAMIC BEHAVIOR OF HEME OXYGENASE Protein dynamics has been largely recognized as an essential determinant of protein function [162–164]. Because the HO catalytic cycle involves multiple changes in coordination and redox states, as well as in the structure of the substrate (heme), while undergoing only minor structural alterations, the polypeptide matrix of HO can be regarded as being inherently dynamic. Furthermore, since HO conserves the fold of the holo-protein when devoid of heme [59,61,165], it seems intuitive that for efficient substrate binding and product (biliverdin) release to occur, the polypeptide must be flexible. Met. Ions Life Sci. 2009, 6, 241–293


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Recent work on bacterial HOs has addressed the issue of protein dynamics and its potential role in enzyme catalysis [120,121]. In what follows, a general overview of the most recent advances related to the dynamic nature of the backbone in HO is presented, followed by a general view of the dynamic behavior that HO exhibits at specific stages of the catalytic cycle, as determined from hydrogen-deuterium (H/D) exchange studies and NMR relaxation measurements conducted with CN, N 3 , and CO-inhibited HO.

4.1. H/D Exchange The propensity of hydrogen atoms of backbone amides to undergo exchange reactions with the solvent in solution is a sensitive indicator of protein flexibility [166,167]. Global probing of protein flexibility by H/D exchange is commonly performed by monitoring the loss of signal intensity in an NMR spectrum as a function of the amount of time that the protein has been in contact with D2O. Values of the exchange rate constants (kex) measured in this fashion are then compared to values of the intrinsic rate constant (kch). The latter, which is a measure of the rate of exchange exhibited by an amide proton in an unstructured polypeptide, is sequence-dependent and can be calculated [168,169]. A commonly used term to express the propensity of amide hydrogens to exchange is the protection factor. Protection factors expressed in logarithmic form (log P ¼ log kex/kch) provide a quantitative estimation of the propensity of a given amide proton to exchange with deuterons in the solvent [168,169]. H/D exchange studies were recently performed with azide- and cyanideinhibited pa-HO [120]. Rates of H/D exchange (kex) were measured for pa-HO-N3 and pa-HO-CN and converted into protection factors. Analysis of the data obtained with each complex are summarized in Figure 11A. To appreciate the information content in this figure it is important to keep in mind that the difference in protection factors, calculated as Dlog P ¼ log PN3 – log PCN, has been plotted per residue. In addition, the horizontal line at Dlog P 0.5 represents the average obtained from summing the absolute value of Dlog P for all residues exhibiting different protection toward exchange. Values of Dlog P ¼ 1.5 correspond to cases where a residue in pa-HO-N3 exchanges with an experimentally measurable kex but the corresponding residue in pa-HO-CN exchanges too fast to allow measurement. This means that although the magnitude of Dlog P cannot be evaluated for these residues, they are clearly located in sections of the protein with the most pronounced differences in propensity to exchange. The data revealed that the majority of residues where there is a difference in propensity to exchange exhibit Dlog P 4 0, which indicates that globally, the backbone amide groups of pa-HO-N3 are distinctively less prone to Met. Ions Life Sci. 2009, 6, 241–293

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Figure 11. (A) Per residue differences in protection factors (Dlog P ¼ log Pazide – log Pcyanide) between pa-HO-N3 and pa-HO-CN. The average difference of all residues is indicated by the horizontal line at Dlog P 0.5. Bars reaching the maximum vertical scale (Dlog P ¼ 1.5) correspond to residues exhibiting exchange rates measurable in pa-HO-N3 but too fast to measure in the pa-HO-CN. (B) View of pa-HO highlighting regions in which pa-HO-N3 has enhanced protection to exchange relative to pa-HO-CN (Dlog P is positive in A). Portions exhibiting 0oDlog Po0.5 are green, 0.5oDlog Po1.5 are yellow and Dlog P ¼ 1.5 are pink. Water molecules are colored blue and are shown as they appear in the X-ray crystal structure (PDB: 1SK7). Adapted from [120].

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exchange than their respective counterparts in pa-HO-CN. These differences, mapped onto the structure of resting state pa-HO (Fig 11B), highlight the most protected residues (Dlog P ¼ 1.5) in pink, residues with Dlog P4average (0.5) in yellow and residues with Dlog Poaverage in green. Remarkable about these findings is the fact that the nature of the distal ligand, the diatomic CN or the triatomic N 3 , appear to exert a large influence on the flexibility of the enzyme, not only in regions of close proximity to the heme, but also in portions of the protein far removed from it. Among residues that are most protected in pa-HO-N3 relative to pa-HOCN (pink and yellow in Figure 11B), those located in helices V, VI, and X and the loop preceding helix X stand out because they are remote from the heme iron and distal ligand. Interestingly, inclusion of the crystallographic structural waters in the model (blue in Figure 11B) strongly suggests the possibility that long range communication between the distal ligand and helices V, VI, and X occurs via these networked structural waters. As has been discussed above, this H-bond network is thought to function in HO as an integral part of a proton delivery machinery used to protonate the nascent FeIII-OO moiety in the process of O2 activation leading to meso-carbon hydroxylation [84,94]. The observations made with H/D exchange experiments, therefore, suggest that proton delivery is not the only function of the network of H-bonded water molecules. Instead, the data implies the tantalizing possibility that an additional function of the structural waters in HO is to provide adaptable interactions between otherwise remote structural elements, thus permitting rapid propagation of conformational changes in the active site with minimum perturbation of secondary structure.

4.2. Microsecond-Millisecond Dynamics NMR spectroscopy has emerged as the technique of choice to gain insights into conformational motions of enzymes in solution. NMR relaxation experiments designed to quantitatively determine the contribution of conformational exchange to the overall transverse relaxation rates of 15N nuclei have been proven particularly powerful in this regard [170,171]. Such an approach was recently applied to study pa-HO-N3, pa-HO-CN, and pa-HO-CO, in order to directly establish whether the axial ligand exerts measurable effects on the dynamic characteristics of the protein backbone [121]. The information derived from these studies, which is complementary to that obtained from H/D exchange measurements, allowed for direct identification of amino acids undergoing ms-ms conformational motions. To achieve this end, the relaxation-compensated Carr-Purcell-Meiboom-Gill Met. Ions Life Sci. 2009, 6, 241–293

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(rc-CPMG) pulse sequence was used to measure the effective transverse 15 N nuclei in pa-HO-N3 and pa-HO-CN. relaxation rates (DReff 2 ) of A comparative analysis of the DReff 2 values obtained from these experiments revealed that, similar to the findings obtained from the H/D exchange studies, CN and N 3 exert a distinct influence on the dynamic behavior of the protein backbone. Again, the most significant differences between the CN and N 3 inhibited enzymes were located in regions that are far removed from the heme iron. It is noteworthy that residues whose relaxation rates were differentially affected by the nature of the axial ligand are involved in hydrogen bonding interactions, via backbone or corresponding side chain, with water molecules in the hydrogen bonding network [121] (Figure 12). This indicates that the differential effects exerted by the ligand on backbone dynamics can be accounted for if direct participation of the hydrogen bonding network is invoked. Consequently, the hydrogen bonding network may be envisioned as a plastic structural motif that allows HO to adapt readily to changes in the reactive (coordination) state of the heme iron. To directly test the involvement of the H-bond network in modulating backbone dynamics in HO, the Arg80Leu mutant of pa-HO (R80L-pa-HO)

Figure 12. Blown-out view of pa-HO (PDB:1sk7) highlighting the hydrogen bonding interactions between water molecules and amino acid side chains undergoing ms-ms backbone conformational motions. Roman numerals are used to identify helices in the structure according to the nomenclature used in the PDB file. Arabic numerals identifying crystallographic waters (blue spheres) are given as assigned in the structure coordinates. Reproduced from [121] with permission of the American Chemical Society, copyright (2007). Met. Ions Life Sci. 2009, 6, 241–293


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was prepared and used to measure per-residue values of DReff 2 . R80 in pa-HO is intimately involved in the hydrogen bonding network of this enzyme (see Figure 12) and is equivalent to D140 in HO-1 (Section 2.4). Replacing this residue for an aliphatic side chain disrupts the hydrogen bonding network and significantly decrease the catalytic activity of pa-HO [121]. Remarkably, a large number of residues in the R80L-pa-HO-N3 mutant undergo ms-ms motions, as is indicated in Figure 13B by the large number of residues with DReff 2 values larger than the average, which is indicated by the segmented horizontal line. This is in stark contrast to the nearly complete ms-ms stillness of the wild-type pa-HO-N3 complex, which is apparent from the few residues with DReff 2 values above the average in Figure 13A. A comparison of data in Figures 13A and 13B, therefore, reveals that residues unaffected by conformational exchange in wild-type pa-HO-N3 become clearly affected by ms-ms motions in R80L-pa-HO-N3. These findings, which were interpreted to be indicative of chaotic motions provoked by the severe disruption of the H-bonding network, highlight the importance

Figure 13. Conformational motions in wild-type pa-HO-N3 (A) and R80L-pa-HON3 (B). Amino acid residues exhibiting DReff 2 values greater than three standard deviations above the average (dashed line) are affected by ms-ms timescale conformational exchange. The widespread occurrence of residues affected by conformational exchange in R80L-pa-HO-N3 relative to wild-type pa-HO-N3 is indicative of the role of the hydrogen bonding network in modulating the dynamic behavior of the protein. Partial reproduction of Figure 4 in [121] by permission of the American Chemical Society, copyright (2007). Met. Ions Life Sci. 2009, 6, 241–293

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of this structural motif in controlling the dynamic characteristics of the protein [121]. Relaxation measurements were also conducted with CO-inhibited pa-HO. – Unlike the N 3 - and CN -inhibited enzymes, pa-HO-CO possesses iron in the +2 oxidation state, which renders it diamagnetic and more amenable for NMR spectroscopic investigations, which allows dynamic characterization of regions near the heme. These investigations showed that residues located in the distal helix, in the region known as the helix kink, are relatively rigid with respect to ms-ms dynamics. It is possible that the dynamic characteristics exhibited by ferrous pa-HO-CO approximate the dynamic state of the pa-HO-O2 complex in the catalytic cycle, prior to formation of the ferric peroxo intermediate. In such scenario, the rigidity of the helix kink can be envisioned to conformationally ‘‘organize’’ the enzyme prior to accepting the second electron and therefore facilitate the formation of the FeIII-O-O intermediate [121]. The effects of CO-inhibition on the ms-ms motions of the pa-HO-R80 mutant have also been studied. Surprisingly, per-residue DReff 2 values measured for R80L-pa-HO-CO are similar to those of equivalent residues in wild-type pa-HO-CO, suggesting conformational stillness of the CO-inhibited mutant in this time scale [121]. A remarkable feature of spectra obtained with R80L-pa-HO-CO, however, is the presence of a noticeably greater number of cross peaks in well-resolved sections of the HSQC spectrum, relative to equivalent sections of spectra acquired with wild-type pa-HO-CO (Figure 14). This suggested that a single cross-peak in the spectrum of wild-type pa-HO-CO is split into two or more cross-peaks in the spectrum of R80L-pa-HO-CO. Sequential assignment of the ‘‘doubled’’ resonances confirmed that single amino acids in R80L-pa-HO-CO produce two well-resolved cross-peaks, indicating the existence of at least two distinct conformations of the mutant in solution. The conformational heterogeneity of the CO-inhibited mutant exchanges slowly (4300 ms) compared to the ms-ms motions of the ferric R80L-pa-HO-N3 complex. These observations demonstrated that the time scale of the conformational disorder brought about by disruption of the H-bond network in the R80L-pa-HO mutant are clearly dependent on both, the nature of the distal axial ligand and the oxidation state of the heme iron [121].

4.3. A Unifying View of Protein Dynamics and Heme Oxygenase Reactivity 1

H NMR spectroscopic investigations conducted with human HO-1 and nm-HO axially coordinated by water, cyanide, and hydroxide in the distal side have established that the hydrogen bond network in these enzymes is Met. Ions Life Sci. 2009, 6, 241–293


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Figure 14. Representative expansions of 1H-15N HSQC spectra of ferrous [U-15N]pa-HO-CO (A) and ferrous [U-15N]-R80L-pa-HO-CO (B). Labels identify crosspeaks according to their sequential assignment. A single backbone amide gives rise to two cross-peaks in spectrum B; indicating the presence of two distinct conformations of R80L-pa-HO-CO in solution. In contrast, equivalent residues in pa-HO-CO give rise to a single cross-peak as shown in spectrum A. Reproduced partially from Figure 10 in [121] by permission of the American Chemical Society, copyright (2007).

structurally robust [62,146,153,172]. This notion was inferred from the low level of deuterium incorporation into amide groups of residues located near the active site after the protein was exposed to D2O for extended periods of time. In addition, results from recent theoretical QM/MM calculations suggest that while robust, the H-bonding network it is prone to undergo reorganization caused by changes in the pattern of H bond to and from water. The robustness of the H bond network was rationalized to exist in order to restrict the orientation and location of the OH radical in the active site prior to hydroxylation of heme [173]. In the context of the experimental dynamic behavior discussed above, it is likely that the ability of the water network to alter its H-bonding pattern is also related to the need that HO has to exert dynamic control on remote regions of the enzyme, as suggested by the NMR relaxation and H/D exchange data described above. Hence, the emerging picture seems to suggest that while maintaining a rigid structure near the active site is important to stabilize reactive intermediates for oxygen activation and heme cleavage, exerting dynamic control over residues located in the protein periphery may be equally essential to facilitate substrate binding, docking with the reductase that provides the electrons needed in the catalytic cycle and product release. Although more experimentation is needed to corroborate this assertion, it seems undeniable that the fold and sequence of HO is exquisitely equipped to sense the changes in redox and coordination state that occur Met. Ions Life Sci. 2009, 6, 241–293

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in the active site at each stage of the catalytic cycle. These changes, in turn, elicit signals that instill in the protein the need to tune the dynamic behavior of the polypeptide, according to the demands imposed by the distinct stages of the catalytic cycle.

5. THE REGIOSELECTIVITY OF HEME HYDROXYLATION 5.1. Hydroxylation of the a-Meso Carbon Leads to Its Release as CO with Subsequent Formation of a-Biliverdin The catalytic activity of HO produces iron, CO, and exclusively one isomer of biliverdin; typically a-biliverdin. This high level of regioselectivity is dictated at the stage of meso carbon hydroxylation because the hydroxylated meso carbon is subsequently released from the macrocycle as CO with the concomitant formation of the corresponding isomer of verdoheme and subsequently, biliverdin (see Figure 1). This section summarizes findings from work aimed at understanding the properties of HO that allow it to convert heme to biliverdin with a high degree of regioselectivity. The crystal structure of human HO-1 revealed that its distal pocket is considerably more polar than that of globins and cytochromes and exhibits several polypeptide backbone-heme contacts (see Figure 4) [53]. The distal helix approaches the heme within 3–4 A˚ along the entire width of the macrocycle and therefore provides steric protection to the b-, g-, and d-meso carbons. This observation strongly suggested that exclusive attack of the unprotected a-meso carbon is controlled by steric steering of the HOO moiety in the FeIII-OOH complex [53]. Corroborating evidence for significant steric control of regioselectivity in heme hydroxylation came from the crystal structure of the azide (FeIII-N3) complex of rat HO-1 [81]. As is clearly illustrated in Figure 15, the coordinated azide molecule is bent (FeIII-N(3)-N(1) angle ¼ 1161) and directed toward the ameso carbon [81]. Interactions between the coordinated N 3 molecule and Gly139, Ser142, Gly143, and Gly144 not only orient the terminal N(1) atom to lie almost on top of the a-meso carbon, but also provide steric protection to the b-, g-, and d-meso carbons [81]. A very similar picture has emerged from the crystallization of the FeII-O2 complex of cd-HO, which demonstrates that the O2 molecule is bent toward the a-meso axis and prevents access to the b-, g-, and d-meso carbons by steric interactions with Gly135, 139, and 140 [174]. These observations also underscore the importance of the Gly residues in the distal helix of HO, Met. Ions Life Sci. 2009, 6, 241–293


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Figure 15. Space filling representation illustrating the interactions between coordinated azide (FeIII-N3) and Gly139, Ser142, Gly143, and Gly144 in rat HO-1, which are responsible for steering the terminal nitrogen (N1) toward the a-meso carbon. Similar steric interactions are expected to orient the HOO ligand to facilitate a-meso hydroxylation. Adapted from reference [81]. Color key: heme (red), coordinated azide (yellow), Gly139 (green), Ser142 (magenta), Gly143 (cyan) and Gly144 (orange).

which serve at least three critical roles: (i) impart flexibility for binding and release of substrate and product, (ii) make close contacts with the heme and coordinated HOO ligand with the purpose of not only steering the latter toward the a-meso carbon but also protecting the remaining meso carbons, and (iii) donate a hydrogen bond to the coordinated O in FeIII-OOH to lower its ligand field strength, thereby promoting the electronic structure that activates the substrate (heme).

5.2. pa-Heme Oxygenase Exhibits Unique d-Regioselectivity Although most of the discussion so far has been in the context of understanding the exquisite a-regioselectivity exhibited by heme oxygenase, it is interesting at this point to discuss the only known HO that oxidizes heme at a carbon other than the a-meso carbon. This discussion is illustrative not only because these investigations contributed significantly to the notion that polypeptide-heme propionate interactions control the regioselectivity of heme oxidation, but also because it illustrates the usefulness of NMR spectroscopy applied to these systems. Heme oxygenase from Pseudomonas aeruginosa, coded by the Pseudomonas iron inducible gene Pig A, was initially reported to oxidize heme to produce mostly b-biliverdin [33]. Met. Ions Life Sci. 2009, 6, 241–293

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Figure 16. (A) The heme seating in pa-HO locates the d-meso carbon where it is susceptible to hydroxylation by the sterically constrained FeIII-OOH moiety. Rotation of the heme by 1801 about the a-g-meso axis places the b-meso carbon where it is susceptible to hydroxylation. (B) The heme in all a-hydroxylating HOs of known structure is rotated B1001 relative to the heme seating in pa-HO, thus placing the a-meso carbon where it is susceptible to hydroxylation by the sterically constrained FeIII-OOH group. Color key: heme (red), proximal histidine (yellow); Phe117 in pa-HO, equivalent to Tyr112 in nm-HO (blue); Asn19 in pa-HO, equivalent to Lys16 in nm-HO (green), Lys34 in pa-HO (magenta). PDB numbers are 1SK7 and iJ77, respectively.

Subsequent studies established that pa-HO oxidizes heme to a mixture of b- (30%) and d-biliverdin (70%) [147]. NMR spectroscopic investigations conducted with the cyanide-inhibited enzyme revealed that the heme in pa-HO is rotated in-plane approximately 1001 relative to the heme in all other a-hydroxylating heme oxygenase enzymes (see Figure 16A). This inplane rotation locates the d-meso carbon within the pa-HO fold in the same place where all a-hydroxylating heme oxygenase enzymes place the a-meso carbon [147] (see Figure 16B). It is also clear that a 1801 rotation of the heme in Figure 16A about the a-g-meso axis places the b-meso carbon in pa-HO where it can be hydroxylated, thus explaining the formation of 30% b-biliverdin [147]. Amino acid sequence alignments in the context of the known structures of heme oxygenase suggested that in-plane rotation of the heme in pa-HO Met. Ions Life Sci. 2009, 6, 241–293


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stems from the absence of hydrogen bonding and electrostatic interactions between the heme propionates and the side chains of Asn19 and Phe117 (see Figure 16A). When these residues in pa-HO were mutated for Lys and Tyr, respectively, in an attempt to introduce the hydrogen bonding and electrostatic interactions typical of heme propionates in a-hydroxylating heme oxygenase enzymes (see Figure 16B), the heme was found to undergo a dynamic in-plane rotation (B1001) [147]. NMR spectroscopic experiments also revealed that the in-plane rotation of the heme exchanges the position of the a-meso carbon in one of the in-plane conformers (heme seating) with the d-meso carbon of the second heme seating. As a result of this dynamic equilibrium the N19K/F117Y double mutant of pa-HO oxidizes heme to a mixture of 55% a-, 35% d-, and 10% b-biliverdin [147]. a-Biliverdin is produced from the heme in-plane conformation (seating) similar to that of a-hydroxylating heme oxygenase (Figure 16B), d-biliverdin is produced from the heme seating characteristic of pa-HO (Figure 16A) and b-biliverdin is produced from a heme orientational isomer obtained by rotating the heme in Figure 16A 1801 about the a-g-meso axis [147]. The advent of the X-ray crystal structure of pa-HO [57] corroborated the unusual seating of the heme in pa-HO and demonstrated that a key interaction between Lys34 and heme propionate-7 (see Figure 16A) stabilizes the heme seating that in pa-HO is conducive to b- and d-hydroxylation [57]. A similar conclusion was reached almost simultaneously from site-directed mutagenesis investigations [175]. The high amplitude in-plane rotation of the heme (B1001), which occurs in a submillisecond time scale, is highly unusual for heme proteins. Typically, steric interactions between heme substituents and side chains lining the heme pocket provide a strong steric friction that maintains the heme in place. Although these interactions are not as pronounced in heme oxygenase, the unusual large-amplitude in-plane rotation of the heme is likely accompanied by polypeptide motions that facilitate the long in-plane excursions of the heme in HO. Although this phenomenon was initially observed only with the N19K/F117Y mutant of pa-HO [147], subsequent work clearly demonstrated that removing key heme propionate-polypeptide interactions leads to large-amplitude in-plane heme disorder and consequently to changes in regioselectivity [147,150,176,177]. The following section summarizes the findings leading to this conclusion.

5.3. Polypeptide-Heme Interactions Control the Regioselectivity of Heme Oxidation The first documented change of regioselectivity brought about by site directed mutagenesis of a heme oxygenase enzyme was observed with the R183D and R183E mutants of rat HO-1, which under coupled oxidation Met. Ions Life Sci. 2009, 6, 241–293

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conditions render a 65:35 mixture of a- and b-biliverdin, respectively [178]. These investigators proposed two plausible mechanisms to explain the change in regioselectivity: (1) Electrostatic repulsion between D183 or E183 and one of the heme propionates, which force the heme to rotate and (2) formation of a new hydrogen bonding network caused by rearrangement of S142 and K179. The mechanism invoking heme rotation was disfavored on the basis that the pattern of paramagnetic heme methyl resonances of the R183 mutants is very similar to that exhibited by wild-type rat HO-1 [178]. This issue was reexamined by studying the corresponding R177D and R177E mutants in heme oxygenase from C. diphtheriae [150]. Replacement of R177 for E or D in cd-HO results in the oxidation of heme to a mixture consisting of B50% of each, a- and d-biliverdin. Detailed NMR spectroscopic analysis of the mutants revealed that formation of d-biliverdin in the mutant stems from electrostatic repulsion of the heme propionates by the side chain of D or E at position 177. Hence, in addition to demonstrating that electrostatic repulsion between the negatively charged E-177 and heme propionates indeed causes heme rotation, the findings from this study contributed to strengthen the notion that the HOO ligand is sterically restrained and oriented toward the a-meso carbon, or whichever meso carbon takes its place upon in-plane heme rotation [147,150]. Subsequent NMR spectroscopic investigations carried out with mutants of HO-1 also demonstrated that placement of negatively charged side chains in close proximity to one of the heme propionates causes B901 in-plane rotation of the heme, which was also accompanied by formation of a mixture of a- and d-biliverdin [177]. To further investigate the idea that key heme propionate-polypeptide interactions promote the appropriate in-plane heme conformation that positions the correct meso carbon where it is unhindered to react with the FeIII-OOH intermediate, a chimeric protein was constructed and investigated spectroscopically [176]. The chimeric protein was prepared by replacing the distal helix of the a-regioselective nm-HO with the distal helix of the d-regioselective pa-HO, as shown schematically in Figure 17. The resultant nm-HO chimera, nm-HOch hereafter, oxidizes heme to produce primarily (95%) a-biliverdin and a very small amount of d-biliverdin (5%). Since the chimeric protein harbors the distal helix (and environment) of the d-biliverdin producing pa-HO, the formation of mostly a-biliverdin clearly eliminates the possibility that the regioselectivity of heme hydroxylation is determined by hydrogen bonding interactions in the distal pocket that steer the hydroperoxide ligand toward a-meso carbon. NMR spectroscopic studies conducted with nm-HOch revealed in-plane disorder of the heme. However, the angle of rotation, although large, is not sufficient to place the d-meso carbon where it can be efficiently attacked by the FeIII-OOH oxidizing species. Thus, the 5% d-biliverdin produced is likely a representation Met. Ions Life Sci. 2009, 6, 241–293


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Figure 17. The chimeric protein, nm-HOch (bottom) was constructed by replacing distal helix residues 107-142 (green) in nm-HO with the corresponding residues (112-147) in pa-HO (purple). The proximal helix is shown in cyan. Adapted from [176].

of the population of molecules where in-plane heme rotation places the d-meso carbon where it is susceptible to attack [176]. To understand the origin of in-plane disorder in the chimeric enzyme it is important to consider that replacement of the distal helix in nm-HOch substitutes Tyr112 in nm-HO (see Figure 18B) with the equivalent residue in pa-HO, Phe117 (see Figure 18A). This eliminates a stabilizing H-bonding interaction between Tyr112 and heme propionate-7 in the nm-HOch protein and ‘‘loosens’’ the heme, which is now able to undergo relatively large inplane rotations. To test this notion the authors eliminated the second hemepropionate-polypeptide interaction in nm-HO by replacing Lys16 with Ala in the nm-HOch to create the nm-HOch-K16A mutant [176]. This protein oxidizes heme to 60% a- and 40% d-biliverdin, a mixture that suggested an increased population of the heme in-plane conformation that places the d-meso carbon where it can be attacked by the FeIII-OOH moiety. This conclusion was supported by NMR spectroscopic analysis of the nm-HOch K16A mutant, where the heme was also found to be in a dynamic equilibrium between two in-plane conformations. One of these conformations makes the a-meso carbon susceptible to hydroxylation, whereas the other in-plane conformation presents the a-meso carbon to the FeIII-hydroxylating group. The information thus far indicated that absence of heme propionatepolypeptide interactions triggers the large in-plane heme disorder seen in the Met. Ions Life Sci. 2009, 6, 241–293

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Figure 18. Crystal structures of pa-HO (A) and nm-HO (B) highlighting the positions of key amino acids in the control of in-plane heme conformation, which in turn controls the regioselectivity of heme oxidation. Note that the in-plane heme orientation of the heme in the two enzymes is related to one another by B901 in-plane rotation. The crystal structures of pa-HO and nm-HO are Protein Data Bank entries 1SK7 and 1J77, respectively. Reproduced from [176] by permission of the American Chemical Society, copyright (2005).

nm-HOch protein. Consequently, if heme propionate-polypeptide interactions are paramount to direct the regioselectivity of heme oxidation by anchoring the heme in an appropriate in-plane conformation within the enzyme, it follows that introducing the key interactions seen in the crystal structure of pa-HO into the chimeric protein should render a d-hydroxylating chimera. Results obtained from probing this idea demonstrated that indeed, replacement of Met31 in nm-HO with the equivalent residue in pa-HO, Lys34 (see Figure 18), to make the nm-HOch K16A/M31K double mutant, results in a recombinant enzyme that oxidizes heme to 95% d-biliverdin. NMR analysis of this enzyme corroborated that the majority of molecules (495%) harbor a heme with an in-plane heme conformation that presents the d-meso carbon to the FeIII-OOH moiety.

5.4. The 1H NMR Spectra of Cyanide-Inhibited Heme Oxygenase as a Diagnostic Tool of Heme Oxidation Regioselectivity As has been pointed out above, bacterial HOs participate in the mining of heme-iron. By comparison, HOs in photosynthetic bacteria are involved in the production of biliverdin, which serves as a precursor in the biosynthesis Met. Ions Life Sci. 2009, 6, 241–293


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of cyanobacterial light-harvesting phycobiliproteins and the photoreceptor phytochromes [179]. Although phytochromes are traditionally known as biliprotein photoreceptors in plants and photosynthetic bacteria, these have been recently discovered in non-photosynthetic bacteria [180]. Unlike plant

Figure 19. Downfield portions of the 1H NMR spectra of nm-HO (A), cd-HO (B), pa-HO (C) and BphO (D). The presence of only one heme methyl resonance (3 methyl, or 3Me) is diagnostic of a-hydroxylating HOs, where the proximal histidineimidazole plane lies approximately parallel to the b-d-meso axis. The same region of the 1H NMR spectrum of pa-HO, where the heme is rotated in-plane B901 relative to the a-hydroxylating HOs shows three methyl resonances (5Me, 1Me, and 8Me). The magnitude of the chemical shifts and the order of these resonances (5Me 4 1Me 4 8Me) in the context of well developed theory [183] indicate that the proximal histidine-imidazole plane in pa-HO lies parallel to the a-g-meso axis. Reproduced from [182] by permission of the American Society for Biochemistry and Molecular Biology, copyright (2004). Met. Ions Life Sci. 2009, 6, 241–293

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and cyanobacterial phytochromes, bacteriophytochromes (BphP) from nonphotosynthetic prokaryotes have been shown to utilize biliverdin as a chromophore [181]. Pseudomonas aeruginosa was among the first bacteria identified to harbor a gene coding for a BphP, which functions as a typical light-regulated histidine kinase. The BphP gene is located in an operon downstream from a BphO gene, which encodes for a putative heme oxygenase enzyme [182]. The product of the BphO gene, BphO, was shown to catalyze the degradation of heme to a-biliverdin, iron, and CO, demonstrating that it is indeed a heme oxygenase [182]. BphO was reconstituted with 13C-labeled heme to assign 1H and 13C resonances from the heme binding site of cyanide inhibited BphO (BphO-CN). Observations made during the study of other cyanide-inhibited bacterial HOs established that the pattern of paramagnetically affected heme methyl resonances in the 1H NMR spectra of cyanide-inhibited HO enzymes (HO-CN) is diagnostic of the heme in-plane orientation, and hence regioselectivity of meso carbon oxidation [147,150]. For instance, Figures 19A and 19B show that only the heme methyl resonance from the major (3Me) and minor (3me) heme orientational isomers in nm-HO-CN [99] and cd-HO-CN [150] is resolved from the diamagnetic region. The 1H NMR spectra of the a-biliverdin producing human and rat HO-1-CN are essentially identical [148,149]. In comparison, the spectrum of pa-HO-CN (Fig. 19C), where the heme is rotated in-plane B1001 relative to the heme in a-biliverdin producing HOs, exhibits three heme methyl resonances (5Me, 1Me and 8Me) resolved from the diamagnetic region [147]. Consequently, the fact that the 1H NMR spectrum of BphO-CN exhibits only the 3Me resonance above 12 ppm (Figure 19D) corroborates that BphO is an a-biliverdin producing HO [182]. Further, the order of the heme methyl resonances in BphO (3Me 4 8Me 4 5Me 4 1Me) indicates that the proximal His-imidazole plane forms an angle j B1331 with respect to the molecular x axis. This means that the proximal His-imidazole plane lies almost parallel to the b-d-meso axis, as is the case for all a-biliverdin producing HOs of known structure (human [53] and rat [54] HO-1, cd-HO [56], and nm-HO [55]). Consequently, it is possible to conclude that the pattern of heme methyl resonances in the 1H NMR spectrum of CN–-inhibited HOs is an effective diagnostic tool of heme oxidation regiochemistry [182].

6. CONCLUSION AND OUTLOOK The catalytic cycle of heme degradation by HO follows a complicated set of reactions that are not yet fully understood. Key to the process is the Met. Ions Life Sci. 2009, 6, 241–293


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generation of a ferric hydroperoxide intermediate in the active site of HO, which is at a crossroad of reactivity because hydroxylation of heme commits the process to heme degradation, whereas formation of compound I is a trademark of monooxygenase and peroxidase enzymes. Hence, the path followed by the ferric hydroperoxide intermediate has enormous repercussions on the biochemistry and biology of O2 activation at heme centers. Despite its importance, we have yet to attain detailed understanding of the determinants that allow HO to bind O2, reduce it to FeIII-OO, deliver a proton to form FeIII-OOH and finally hydroxylate the heme, while avoiding being inhibited by the CO released from the process. A very large effort encompassing chemical, biochemical and theoretical approaches, as well as biophysical techniques such as X-ray crystallography, EPR, NMR, resonance Raman and other spectroscopies, have shed significant light on the process. Nevertheless a significant portion of the path remains uncharted. Among the challenges to overcome in the future is the study of the electronic structure of the highly reactive ferric peroxide and ferric hydroperoxide intermediates at ambient temperatures, because investigations with model heme complexes suggest that their electronic structures at physiologically relevant temperatures are distinct from those observed at the cryogenic temperatures used to trap and study them. In addition, recent investigations suggest that dynamic motions involving the hydrogen bonding network in the distal site of HO are significantly affected by the nature of the distal ligand and by the oxidation state of the heme-iron. Moreover, disruption of the hydrogen bonding network creates global conformational disorder of the enzyme in the ms-ms time scale, which is accompanied by a decrease in reactivity. Thus, in order to fully understand the control that enzyme dynamics exerts on reactivity of heme degradation it will be important to conduct studies aimed at comparing the polypeptide motions in distinct coordination and oxidation states with specific rates of ligand binding and O2 activation. Although challenging, investigations of this nature would not only generate important information to further understand the mechanism of O2 activation by HO including its inhibition by CO, but would also represent an unprecedented description of dynamic-reactivity relationships in heme containing proteins and enzymes.

ACKNOWLEDGMENTS The author’s research reported in this manuscript was carried out with support from grants from the National Institutes of Health (GM 50503) and the National Science Foundation (MCB-0446326). Met. Ions Life Sci. 2009, 6, 241–293


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ABBREVIATIONS BphP cd CPR cyt P450 ENDOR EPR FAD FMN FPR FRET H/D Hb Hb-CO h-HO-1 HO HSQC Mb MM NADPH nm nm-HOch NMR pa Por QM rc-CPMG r-HO-1 SPR

bacteriophytochrome Corynebacterium diphteriae cytrochrome P450 reductase cytochrome P450 electron nuclear double resonance electron paramagnetic resonance flavin adenine dinucleotide flavin mononucleotide NADPH-dependent ferredoxin reductase fluorescence resonance energy transfer hydrogen-deuterium hemoglobin carboxyhemoglobin human heme oxygenase-1 heme oxygenase heteronuclear single quantum coherence spectroscopy myoglobin molecular mechanics nicotinamide dinucleotide phosphate reduced Neisseria meningitidis chimeric nm-HO nuclear magnetic resonance Pseudomonas aeruginosa porphyrin quantum mechanics relaxation-compensated Carr-Purcell-Meiboom-Gill rat heme oxygenase-1 surface plasmon resonance

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9 Copper-Carbon Bonds in Mechanistic and Structural Probing of Proteins as well as in Situations where Copper is a Catalytic or Receptor Site Heather R. Lucas and Kenneth D. Karlin Department of Chemistry, The Johns Hopkins University, 3400 N. Charles Street, Baltimore, MD 21218, USA

ABSTRACT 296 1. INTRODUCTION 297 2. BINUCLEAR COPPER PROTEINS 298 2.1. Coupled Binuclear Copper Proteins 298 2.1.1. Carbon Monoxide versus Dioxygen Binding to Copper(I) 300 2.1.2. Relationship of Overall Structure to Protein Function 302 2.1.3. Trends in Spectroscopic Properties of CarbonmonoxyHemocyanin and Tyrosinase 304 2.2. Noncoupled Binuclear Copper Proteins 308 2.2.1. Static Structure of the Catalytic Core of Peptidylglycine a-Hydroxylating Monooxygenase as Determined by X-Ray Crystallography 309 2.2.2. Active Site Probing of the Catalytic Core of Peptidylglycine a-Hydroxylating Monooxygenase and Dopamine b-Monooxygenase through Carbon Monoxide Coordination 311


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2.2.3. Coordination of Isocyanide to CuM and CuH in the Catalytic Core of Peptidylglycine a-Hydroxylating Monooxygenase and Dopamine b-Monooxygenase 3. HETEROBIMETALLIC COPPER-CONTAINING ENZYMES 3.1. Heme-Copper Oxidases 3.1.1. Structural and Mechanistic Probing through Photochemical Methods 3.1.2. Proton Translocation Pathways 3.1.3. Mixed-Valent Heme-Copper Oxidases and Electron Transfer 3.1.4. Interaction of Cyanide 3.1.5. Nitric Oxide Reduction Capabilities of Cytochrome c Oxidase 3.2. Cu-Zn Superoxide Dismutase 3.3. Molybdenum-Copper Carbon Monoxide Dehydrogenase 4. NON-BLUE COPPER OXIDASES 4.1. Copper Amine Oxidase 4.2. Galactose Oxidase 5. BLUE, GREEN, AND PURPLE COPPER PROTEINS 5.1. Blue Electron Transfer Proteins 5.2. Multi-Copper Oxidases 5.3. Copper Enzymes in Denitrification 5.3.1. Nitrite Reductase 5.3.2. Nitrous Oxide Reductase 6. COPPER(I) RECOGNITION SITES OR RECEPTORS 6.1. Bacterial Copper Chaperone CusF 6.2. Copper-Ethylene Receptor 6.3. Copper Ion in an Olfactory Receptor Site? 7. MISCELLANEOUS 7.1. Bleomycin 7.2. Copper-Alkyl Complexes from Biologically Derived Carbon Radicals 8. GENERAL CONCLUSIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

315 317 317 318 323 324 327 329 330 332 334 334 337 337 337 338 340 340 342 344 344 345 345 346 346 349 349 350 351 352

ABSTRACT: While copper-carbon bonds are well appreciated in organometallic synthetic chemistry, such occurrences are less known in biological settings. By far, the greatest incidence of copper-carbon moieties is in bioinorganic research aimed at probing copper protein active site structure and mechanism; for example, carbon monoxide (CO) binding as a surrogate for O2. Using infrared (IR) spectroscopy, CO coordination to cuprous sites has proven to be an extremely useful tool for determining active site copper ligation (e.g., donor atom number and type). The coupled (hemocyanin,

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tyrosinase, catechol oxidase) and non-coupled (peptidylglycine a-hydroxylating monooxygenase, dopamine b-monooxygenase) binuclear copper proteins as well as the hemecopper oxidases (HCOs) have been studied extensively via this method. In addition, environmental changes within the vicinity of the active site have been determined based on shifts in the CO stretching frequencies, such as for copper amine oxidases, nitrite reductases and again in the binuclear proteins and HCOs. In many situations, spectroscopic monitoring has provided kinetic and thermodynamic data on CuI-CO formation and CO dissociation from copper(I); recently, processes occurring on a femtosecond timescale have been reported. Copper-cyano moieties have also been useful for obtaining insights into the active site structure and mechanisms of copper-zinc superoxide dismutase, azurin, nitrous oxide reductase, and multi-copper oxidases. Cyanide is a good ligand for both copper(I) and copper(II), therefore multiple physical-spectroscopic techniques can be applied. A more obvious occurrence of a ‘‘Cu-C’’ moiety was recently described for a CO dehydrogenase which contains a novel molybdenum-copper catalytic site. A bacterial copper chaperone (CusF) was recently established to have a novel d-p interaction comprised of copper(I) with the arene containing side-chain of a tryptophan amino acid residue. Meanwhile, good evidence exists that a plant receptor site (ETR1) utilizes copper(I) to sense ethylene, a growth hormone. A copper olfactory receptor has also been suggested. All of the above mentioned occurrences or uses of carbon-containing substrates and/or probes are reviewed and discussed within the framework of copper proteins and other relevant systems. KEYWORDS: carbon monoxide copper proteins copper receptors cyanide ethylene and cation-p-interactions heterobimetallic (Fe, Zn, Mo) units isocyanide

1. INTRODUCTION Even for most biochemical or inorganic researchers focusing on metals in biology, the bioinorganic chemistry of the copper-carbon bond will probably not easily come to mind. For example, the vast majority of proteins or enzymes with copper ion containing active sites deal either with electron transfer (e.g., azurin or plastocyanin), nitrogen oxide processing (for nitrite reduction to nitrogen monoxide (dNO) or nitrous oxide (N2O) reduction to N2), or dioxygen processing. The latter include the O2-carrier hemocyanins, coupled or uncoupled dicopper monoxygenases, copper oxidases which couple substrate oxidation/dehydrogenation to O2-reduction to H2O2 or water and others. In none of these very many cases are there carbon based active-site ligands; by contrast, carbon monoxide (CO) and cyanide (CN–) are natural metal-ligands in iron or iron-nickel hydrogenases (see Chapters 5, 6, and 7 in this volume). And for NOx and O2-processing proteins, the copper binding substrates are nitrogen oxide- or dioxygen-derived small molecules. It is the purpose of this review to survey the incidence of carbon-based metal ligands in copper bioinorganic chemistry. In fact, there is a substantial literature such that we will only highlight the field, while trying to provide details in many of the cases. The major area of occurrence of Cu-C bonds is in the use of chemical and/or spectroscopic probing of copper protein active Met. Ions Life Sci. 2009, 6, 295–361

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sites. Such applications go back as far as 1919, when Craifaleanu observed that bubbling of CO through solutions of a highly colored hemocyanin (oxyform) led to a change to colorless [1]. This was again observed in 1922 by Dhere and Schneider, but they showed that hemocyanin (Hc) previously exposed to CO would recolor when exposed to air. More detailed insights obtained by Root in 1934 [1] revealed that both CO and O2 bind to hemocyanin in a two-copper to one small molecule stoichiometry, and that in sharp contrast to the behavior of hemoglobin, CO binds less strongly (Keq about 20 times smaller) to copper in Hc than does O2. Thus, just as has been extensively employed for the interrogation of heme protein iron centers, CO, CN– and even isocyanides (RNC) have been and are used for copper proteins. This is particularly true for the investigation of copper(I), the reduced redox partner of copper(II), in proteins processing nitrogen oxides and dioxygen. As is well known from inorganicorganometallic chemistry, these three carbon-based ligands are good p-acceptors, thus excellent ligands for low-valent copper(I). Cuprous ion is also well known to ligate to olefins and even arenes, and as such we will draw attention to copper proteins where Cu(I)-R 0 (R 0 ¼ olefin or arene) interactions are intimately involved. As cuprous ion is a d10 metal ion, there are few spectroscopic handles; for example, UV-vis and EPR spectroscopies do not apply. However, IR signatures for CO, CN, and RNC tell a great deal about local environment. As an anionic ligand, cyanide is also a very good ligand for cupric ion, and as will be seen, various spectroscopic methods can then be utilized to interrogate the ligand field about Cu(II), thus obtaining insights into ligation at copper protein active sites, these being new entities in coordination chemistry. As for all of biological chemistry, structure and function are integrally intertwined. To fully comprehend enzyme reactivity and mechanisms, active site structural insights for both Cu(I) and Cu(II) are key. In fact, in one biological example, CO is produced as a product of the copper dioxygenase protein quercetinase, although it does not appear that a Cu-CO interaction ever occurs. However, very recent and exciting findings reveal that copper(I) binds carbon monoxide as the enzyme substrate in molybdenumcopper CO dehydrogenase (MoCuCODH), involving the oxidation of CO to CO2. This system will be discussed in some detail (see Section 3.3).

2. BINUCLEAR COPPER PROTEINS 2.1. Coupled Binuclear Copper Proteins Hemocyanin, tyrosinase (Tyr), and catechol oxidase make up a class of proteins that have undergone study for over a century [1–3]. Each protein Met. Ions Life Sci. 2009, 6, 295–361

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Figure 1. X-ray crystal structures of deoxy-hemocyanin (left) and oxy-hemocyanin (right) from Limulus II (horseshoe crab). Adapted from [4].

has a binuclear active site consisting of two copper centers (labeled CuA and CuB), independently coordinated by three imidazole histidines as shown in Figure 1. The close proximity (2.9–4.6 A˚) of CuA and CuB enables fast electron transfer (ET) for the two electron reduction of dioxygen by deoxy-Hc to form oxy-Hc, best described as a side-on bound m-Z2:Z2peroxo dicopper(II) species [4,5]; there has never been any evidence for an initially formed single-copper ion adduct, Cu-(O2). For oxy-Hc and other met (i.e., oxidized) enzyme forms, a strong magnetic interaction (2J Z 1200 cm1, H ¼ 2JS1 S2) exists between the CuA and CuB ions, which led to their designation by Solomon and coworkers as Type III coupled binuclear proteins [6]. Due to their contradictory functions, the nature of the related coupled binuclear proteins have baffled researchers since their discovery. Hc is the dioxygen transport protein in arthropods and mollusks, analogous to hemoglobin in vertebrates. Tyr is found in all aerobic organisms and is responsible for catalyzing the hydroxylation of monophenols to orthodiphenols and the subsequent two-electron oxidation to ortho-quinones. As illustrated in Scheme 1, the first monoxygenase-type process is referred to as phenolase or cresolase activity and the second oxidase-type process is referred to as catecholase activity. Catechol oxidase is also involved in the production of ortho-quinones but lacks the cresolase ability like that of Tyr. The production of quinone product initiates the synthesis of melanin, which

Scheme 1. Met. Ions Life Sci. 2009, 6, 295–361

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is responsible for the browning processes found in fruits, vegetables, and the skin of mammals [7]. Catechol oxidase is found only in plants and aids in their protection from pathogens or insects.

2.1.1. Carbon Monoxide versus Dioxygen Binding to Copper(I) As mentioned, early work by Root established that only one small molecule, CO or O2, binds per dicopper active site in deoxy-Hc [1], and it binds CO with a much lower affinity than O2. Through isotopic 13C and 18O studies, Fager and Alben confirmed that carbon monoxide coordinates in a terminal fashion to a single copper site rather than in a bridged coordination mode between both copper sites, like for O2 [8]. Extensions by van der Deen and Hoving later confirmed that CO linkage to copper was through the carbon atom [9]. Differences in the observed coordination abilities of the CuA and CuB sites has led to a plethora of research focusing on the chemical nature of the dicopper active site. Bourne and coworkers measured the kinetics and equilibrium binding constants of CO and O2 establishing the cooperative nature in which the multi-subunit Hc’s bind O2, but not carbon monoxide [10]. In establishing fundamental aspects of CO and O2 binding to copper(I) centers, recent studies by Karlin and coworkers have led to the determination of CO binding kinetics and equilibrium parameters for a series of synthetic mononuclear copper(I) compounds possessing tripodal tetradentate ligands [11]. The study of mononuclear copper(I) carbonyl species is very applicable for comparison to Hc since the CO coordination in Hc-CO involves only one copper ion (see Figure 2). As shown in Table 1, the CO binding constant (KCO) for [CuI(tmpa)(Solv)]1 is similar to that for deoxy-Hc [10,11]. The model compound data was further supplemented by transient absorbance laser flash photolysis experiments. By monitoring CO re-binding following photodissociation from [CuI(tmpa)(CO)]1, the CO association rates (kCO)

Figure 2. Schematic of carbonmonoxy-hemocyanin (left) and a mononuclear synthetic carbonyl complex, [CuI(tmpa)(CO)]1 (right). Met. Ions Life Sci. 2009, 6, 295–361

15.4 0.38 (0.74–117) 104 (2.9–48) 105 (2.6–5.4) 105 1.8 105

1.25 105 220 2.6 107 4.6 108 (2.7–11.2) 103

220 103

[CuI(H-tmpa)(THF)]1 [CuI(H-tmpa)(CH3CN)]1 Myoglobin (human Mb) Hemoglobin (human Hb) Limulus polyphemus (arthropodal Hc) Busycon carica (molluskan Hc)

KO2 (M1)

KCO (M1)

Compound or Protein

7.7 105

1.9 109 5.9 107 7.6 105 4.6 106 (2–4.3) 105

kCO (M1 s1)

(1.1–2.2) 106

1.3 109 5.8 107 (1.4–25) 107 (2.9–22) 107 (1.3–1.9) 106

kO2 (M1 s1)

3–4

1.54 104 2.68 105 0.022 0.009 38–75

kCO (s1)

6.5–11.5

1.3 108 1.5 108 22 13.1 2.4–7.5

kO2 (s1)

Table 1. Comparison of O2 and CO rate and binding constants at 298 K for [CuI(H-tmpa)(Solvent)]1 [11], arthropodal and molluskan hemocyanin [10], hemoglobin, and myoglobin [12].

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Table 2. Comparison of CO and O2 equilibrium binding constants for arthropodal and molluskan hemocyanins [14]. Arthropodal and Molluskan Hemocyanins

KCO (M1)

KO2 (M1)

Panulirus interruptus (arthropodal Hc) Limulus polyphemus (arthropodal Hc) Busycon canaliculatum (molluskan Hc) Octopus vulgaris (molluskan Hc) Helix pomatia (molluskan Hc)

2.9 103 (2.4–7.6) 103 2.1 105 3.3 105 (0.8–2.5) 105

6.4 104 (0.35–60) 105 4.3 104 1.1 104 (1.3–6.1) 104

were measured. Further, dissociation rate constants (kCO) were calculated based on the relationship, KCO ¼ kCO/kCO. Additionally Karlin and coworkers have measured O2-binding kinetics (kO2, kO2) and equilibrium constants (KO2) for [CuI(tmpa)(Solv)]1 by taking advantage of the competitive nature of CO and O2 following photodissociation of CO from [CuI(tmpa)(CO)]1 [13] (see Table 1). Similar to hemoglobin and myoglobin, the kO2 values for [CuI(tmpa)(Solv)]1 were higher than the kCO values. However in hemocyanin, the kCO value was greater than the kO2 value resulting in a lower KCO value in comparison to the KO2 value [10]. Arthropodal hemocyanins are comprised of one to eight hexamers with one dicopper binding site per hexamer subunit. In contrast, the functional macromolecules of molluskan species have seven or eight binding sites per decamer subunit [15]. Melzner et al. have suggested such differences to an evolutionary response to the natural habitat or a developed thermal tolerance due to environmental conditions [16]. Brunori et al. determined the CO and O2 coordination affinities for various arthropodal and molluskan species of hemocyanin [14] (see Table 2). Many have attributed the differences to variations in quaternary structures or protein folds that lead to diverse levels of cooperativity upon binding CO and/or O2. Methods for modeling the cooperative nature of the protein active site have been developed, for example the Monod-Wyman-Changeux (MWC) model and its hierarchial extension, the Nested MWC model [15,17]. However, a full description is outside the scope of this article and therefore will not be covered. Instead, functional relationships will be discussed primarily based on the primary structure and second shell ligands.

2.1.2. Relationship of Overall Structure to Protein Function Crystallographic and biochemical studies point to the conclusion that functional differences amongst the coupled binuclear proteins result from Met. Ions Life Sci. 2009, 6, 295–361

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variations in the binding pockets that alter the accessibility of the active sites to substrates [4,17–24]. The tyrosinase and catechol oxidase active sites are positioned near the molecular surface, which ensures that substrates can access the catalytic center [20,21]. However, the active site of hemocyanin is positioned deep within the protein domains [17]. In addition, amino acid residues protruding from different folding domains of hemocyanin further shield the dicopper center. For example, a leucine residue (Leu2830) extends into the protein pocket of Octopus dofleini Hc and a phenylalanine residue (Phe49) is present in Limulus polyphemus Hc [4,18,19]. As a result, only small molecules such as carbon monoxide or dioxygen can coordinate, leading to the protein physiological O2 transporter role. In the recent X-ray crystal structure of Streptomyces tyrosinase solved by Matoba and coworkers [21], a large vacant space was present above the dicopper active site. The vacancy was filled by a tyrosine moiety derived from a ‘‘caddie protein’’ used to obtain crystals; the phenol group was found relatively close to the metal center [21]. In addition, the folding motifs of the protein domain allow for a high degree of flexibility upon redox changes. For example, the distance between the two copper centers is B4.1 A˚ in the fully reduced forms, but decreases to B3.6 A˚ in dicopper(II) forms. The flexibility is likely important for accommodating the bidentate catechol bridging coordination mode (Figure 3) that forms following the phenolase activity exhibited by tyrosinase (see the recent review by Itoh and Fukuzumi for details concerning the hydroxylation mechanism [2]). In the X-ray crystal structure of Ipomoea catechol oxidase (IbCO) from sweet potato, one of the histidine residues (His109) bound to CuA is found to be covalently linked through a thioether bridge to a cysteine residue (Cys92) [20]. The Cys-His crosslink would perhaps put structural restraints on the CuA center leading to greater active site specificity for catechol oxidase in comparison to tyrosinase. In addition, a phenylalanine (Phe261) side chain is

Figure 3. Key intermediates during the different reaction cycles of tyrosinase and catechol oxidase (see text for more detail). Adapted from [25]. Met. Ions Life Sci. 2009, 6, 295–361

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positioned above the CuA center. As a result, phenolic substrates are hindered from CuA and monodentate substrate coordination to CuB is favored. According to a recent computational study by Gue¨ll and Siegbahn, the catecholase cycle (two-electron oxidation of diphenols) would require a monodentate substrate coordination, like that shown in Figure 3 [25]. Therefore, it is likely that only catecholase activity is exhibited by catechol oxidase because the bidentate coordination mode that would follow phenolate hydroxylation is prevented. As proposed by Matoba and coworkers, phenolase and/or catecholase activity may be based on the flexibility or restraints placed on the CuA center [21]. Similar to catechol oxidase, a thioether Cys-His bond that stabilizes the CuA site is observed in the X-ray structure of Octopus Hc [19]. A direct correlation between the function of the protein and the presence of a Cys-His crosslink has not been made because inconsistencies exist. For example, sequence comparisons amongst the binuclear copper proteins have suggested a thioether bridge exists in the active site of most molluscan hemocyanins, such as in Octopus Hc described above, as well as in Neurospora and Agaricus tyrosinases [7,22,26]. However, the Cys-His crosslink is absent in arthropodal hemocyanins and some mammalian tyrosinases [19,27]. Recent work by Decker and coworkers has indicated that hemocyanin can exhibit phenoloxidase or catechol oxidase activity through addition of allosteric effectors under certain conditions [28–30]. Terwilliger has suggested this functional switch as an immune response due to bacterial invasion or extreme changes in environmental conditions [31]. Regardless of these evolutionary implications, variations in the hydrophobic protein pocket likely guide the different active site functions of the coupled binuclear copper proteins.

2.1.3. Trends in Spectroscopic Properties of CarbonmonoxyHemocyanin and Tyrosinase The general overview given above, on the different structural features of the binuclear copper proteins, is important for understanding the spectroscopic and emissive properties of the carbonmonoxy-Hc and tyrosinase species. As will be seen in the discussions to follow, these investigations have not only led to new descriptions and photophysics for novel chemical entities, but have generated further and often detailed descriptions of the protein actives sites and the subtle variations which occur in nature. From early work by Fager and Alben, structural differences were evident amongst the carbon monoxide complexes of hemocyanin from arthropods and mollusks, even before their characterization by X-ray crystallography [8]. For example, the CO stretching frequencies (nCO) of the two mollusk Met. Ions Life Sci. 2009, 6, 295–361

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species studied from Logigo pealii and keyhole limpet (Megathura crenulata) were nCO ¼ 2063 cm1 and nCO ¼ 2062 cm1, respectively. The nCO values were on average nCO ¼ 2043 cm1 for the various arthropod species studied such as Cancer crab (C. irroratus) and both Camberus and Orconectes crayfish (C. robustus, C. loevis, O. rusticus, and O. sanborni). Since X-ray crystal structures have now been solved for other mollusk and arthropod species, it seems reasonable to suggest that the difference in nCO values relates to the nature of the immediate environment around the CuI-CO moiety and even to second-shell ligands of the protein active site. As mentioned above, most molluskan hemocyanins likely possess the Cys-His crosslink, and we suggest that the higher nCO values observed in mollusks are likely due to the difference in donor ability of the Ne-histidines as a result of the Cys-His crosslink. Further, as discussed above with respect to the differences in importance placed upon CuA versus CuB in different proteins or enzymes, shifts in the CO stretching frequencies of different dinuclear copper protein active sites may point to differing preferences for CuA versus CuB substrate coordination. How changes in ligand electron donating ability affect copper(I) reactivity has long been investigated through synthetic model systems [32]. For example, Karlin and coworkers utilized a tridentate N-donor ligand system referred to as R-PYAN (N-[2-(4-R-pyridin-2-yl]-ethyl)-N,N 0 ,N 0 -trimethylpropane-1,3-diamine; R ¼ NMe2, OMe, H, and Cl) to examine variations in copper(I)-dioxygen reactivity [33]. As a probe and model for observed reactions with O2, carbon monoxide adducts of the copper(I)-ligand species were synthesized and the shifts in the CO stretching frequencies were monitored by IR spectroscopy. As the electron-donating ability of the para substituent on the pyridyl donor increases, the nCO value attributed to [CuI(R-PYAN)(CO)]B(C6F5)4 decreases. In addition, the shifts of the nCO values correlate with their CuII/I redox potentials, as illustrated in Figure 4. Therefore, in relation to the hemocyanin species investigated by Fagen and Alben, one may suggest that the copper at the substrate binding sites in

Figure 4. Trends in reduction potentials (E1/2) and CO stretching frequencies (nCO) upon changing the ligand (R-PYAN) electron-donating ability in a synthetic copper(I) complex model system. Adapted from [33]. Met. Ions Life Sci. 2009, 6, 295–361

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arthropods are more strongly bound by histidine in comparison to those in molluscan species [8]. The effect that the thioether bridge would have on the donor ability of the CuA histidine residue has yet to be examined by synthetic model chemists. The luminescence properties of several hemocyanins (molluskan and arthropodal species) have also been extensively studied, in order to further understand the chemical nature of the dinuclear active site [34–38]. The fluorescence profile of the fully reduced binuclear proteins exhibit an emission centered around 335–350 nm arising from energy transfer from the aromatic amino acid residues to the dicopper center. Upon saturation with CO to produce carbonmonoxy-Hc (HcCO) and excitation in the ultraviolet region (280–320 nm), the 350 nm emission is moderately quenched and a new strong emission band arises with a maximum in the 540–560 nm region. This emissive process has a measured lifetime in the typical range of 60–85 ms with quantum yields in the range of f ¼ 0.2–0.4 that has been interpreted as energy transfer from a singlet MLCT band to a low lying triplet state, thus a phosphorescence phenomenon [39]. Similar to other arthropodal hemocyanin species, carbonmonoxy-hemocyanin from horseshoe crab (Limulus polyphemus) also exhibits an emission centered at lem ¼ 560 nm, however, with a much longer lifetime of t ¼ 115 ms. In addition, Limulus HcCO has a nCO value of 2054 cm1 that falls in the middle of the nCO range described above. The likely cause is the presence of a phenylalanine residue (Phe49) positioned above the CuA site [4,18]. In a coordination complex of copper(I) with benzo[h]quinoline ligands capable of p-stacking Thummel and coworkers observed long-lived copper MLCT excited states [40]. A similar phenylalanine residue (Phe261) is positioned above the CuA center of catechol oxidase (see Figure 5C). Therefore, a similar longlived emission spectrum may result upon excitation, however the luminescent properties of carbonmonoxy-catechol oxidase have yet to be investigated. As perhaps is relevant, various copper synthetic model systems have emerged which in fact show how cuprous ion can interact with aromatic psystems. For example, Itoh and coworkers examined copper(I) complexes supported by pyridylalkylamine tridentate ligands with phenylethylene and phenylmethylene appendages which reveal the presence of copper(I)-arene p interactions [41–44]. The strength of the Z2-type copper-arene interaction was altered through the input of different phenylethylene para-substituents, see Figure 5A [42]. Enhanced d-p interaction as a result of increased electron density from the para-substituents also diminished the reactivity of the ligand-copper(I) species toward dioxygen. Also, N-alkyl substituents designated by ‘‘R’’ in Figure 5A were found to hinder the free rotation of the phenylethylene sidearm, further enhancing the d-p interaction [42,44]. No copper-arene interaction resulted when a shorter linker than the aforementioned species was utilized as indicated by the ability of the copper(I) Met. Ions Life Sci. 2009, 6, 295–361

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Figure 5. Comparison of copper(I)-arene p-interactions in synthetic (A, B) and natural systems (C). Modified from [42], [44], and [45]. See text for further discussion.

complex to form a carbonyl adduct (Figure 5B) [44]. The luminescent properties of the ligand-copper(I) species were not investigated. Variable temperature work by Kanagy et al. contributed to the assignment of the emission maximum of most coupled binuclear proteins being designated as a MLCT excited state [36]. As described by Sorrell et al., histidine ligands have low-lying orbitals capable of producing low-energy charge transfer excited states [46,47]. Similar to synthetic model systems, the emission maximum (lem) of most coupled binuclear copper proteins red shift upon lowering the temperature from 298 K to 77 K [36]. Kanagy et al. also showed that the luminescence properties of fungal tyrosinase from Neurospora crassa were very similar to known emission profiles of hemocyanins. However, the emission of Agaricus bisporus tyrosinase from white mushroom slightly blue shifted upon lowering the temperature and as a result the process was believed to derive from a metal-centered excited state [36]. Interestingly, the presence of both copper(I) centers in the binuclear active site is essential for emission even though CO is known to bind to only one site of the dinuclear center. Finazzi-Agro` et al. discovered that the luminescent state of HcCO is absent in the half-met [CuICO..CuII] and half-apo [CuICO..( )] derivatives [37]. Similarly, Jackman, Hajnal, and Lerch determined that replacement of one of the CuA histidine residues from Streptomyces glaucescens tyrosinase with a glutamine or asparagine residue abolished the luminescence exhibited by the carbonmonoxy species [38]. However, mutations at the CuB site had less of an effect suggesting CuA as the substrate coordination site. Other perturbations to the enzyme such as addition of substrate analogs and changes in pH have less of an effect on the luminescence [36,37]. Such a lack of response led to the suggestion that the active moiety is rather shielded or inaccessible, as was later proven by X-ray crystallography (see above). Met. Ions Life Sci. 2009, 6, 295–361

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Furthermore, addition of dioxygen to the copper(I) carbonyl species completely quenches emission at lem ¼ 550 nm and in turn gives rise to the absorbance signature for the side-on m-Z2:Z2-peroxo species around lmax ¼ 340 nm. As stated above, KO2 values are greater than the KCO values likely due to the cooperative nature of the proteins. Brunori and coworkers investigated the different displacement patterns of O2 and CO from the active site of Helix pomatia b-hemocyanin [48]. Addition of small amounts of CN– in the presence of both O2 and CO resulted in a mixed ligand species [CuICO..CuIICN], as a result of O2 displacement by CN–. The simultaneous presence of CO and CN– bound to the active site was supported by an increase in emission intensity and a concomitant decrease of the O2 absorption band. Solomon and coworkers also suggested the existence of a mixed-ligand complex occurring in Busycon canaliculatum half-met hemocyanin [49]. Formation of a mixed ligand species further supports the presence of a highly tuned active site.

2.2. Noncoupled Binuclear Copper Proteins Dopamine b-monooxygenase (DbM) and peptidylglycine a-hydroxylating monooxygenase (PHM) are binuclear copper proteins, however, with uncoupled mononuclear active sites, i.e., they are well separated [50,51]. Both monoxygenases are fascinating in that they activate dioxygen at a single copper site (CuM CuB) for insertion into C-H bonds of prohormones (see Scheme 2). More specifically, DbM is responsible for catalyzing the hydroxylation of phenylethylamines at the benzylic position, such as dopamine to produce norepinephrine. Similarly, PHM catalyzes the


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hydroxylation of peptidylglycine moieties, the first step towards C-terminus peptide amidation; such product peptides are essential for proper control of cellular function. DbM and PHM exhibit a high degree of sequence homology and react towards substrates with similar chemical mechanisms [50,51].

2.2.1. Static Structure of the Catalytic Core of Peptidylglycine a-Hydroxylating Monooxygenase as Determined by X-Ray Crystallography The X-ray crystal structure of a truncated form of PHM, termed PHMcc, i.e., the PHM catalytic core, was reported by Amzel and coworkers for both the oxidized and ascorbate reduced forms of the enzyme [52]. PHMcc consists of only protein domains necessary for monooxygenase activity. As shown in Figure 6, the two copper domains (CuM, CuH) are separated by 11 A˚ [52]. The first domain, which encompasses the catalytic active site (CuM CuB), is in the oxidized state bound in a distorted tetrahedral geometry by two Ne-histidine residues (His242, His244), a methionine sulfur (Met314), and a coordinated water molecule. The second domain is comprised of the electron storage/transfer site copper ion (CuH CuA) which is coordinated by three Nd-histidine residues (His107, His108, and His172) in a T-shaped geometry. Little variation exists in the static X-ray structures upon changes in CuII/I redox state, however other spectroscopic techniques have suggested that indeed there are large structural changes [53].

Figure 6. X-ray crystal structure of PHM displaying a peptidyl substrate near the CuM site. Adapted from [54]. Met. Ions Life Sci. 2009, 6, 295–361

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The more recently reported structure by Amzel and coworkers of a copper-dioxygen precatalytic PHMcc analogue shows that dioxygen binds to CuM as an Z1 (end-on) superoxide moiety (O 2 : one-electron reduced dioxygen) (see Figure 7) [51]. Klinman and coworkers had for many years suggested a hydroperoxo-Cu(II) CuM moiety as the species that leads to the hydroxylation chemistry observed. However, the more recently proposed mechanism involves a copper(II) superoxide moiety, i.e., the initial dioxygen-copper(I) adduct, as the active hydrogen atom abstractor in the catalytic mechanism of PHM and DbM [50,55]. DFT calculations from Chen and Solomon also led to such conclusions; however other key intermediates have also been proposed [56]. For example, a contradictory mechanism that involved ‘superoxide tunneling’ was at one time proposed by Jaron and Blackburn [57,58] due to the unexpected reactivity of the CuH center of PHMcc with CO (see below for further details). Stabilization of copper(II)-superoxide species for detailed chemical and structural analysis is rare for both synthetic and natural systems. Two synthetically derived copper(II) Z1-superoxide complexes were recently described [59–61] (see Figure 7). Schindler and coworkers crystallographically characterized [CuII(TMG3tren)(Z1-O2)]1 (TMG3tren ¼ tris(2-(N-tetramethylguanidyl)ethyl)amine), which upon addition of an H-atom donor can activate C-H bonds through O-atom insertion [59,60]. 1 (NMe2A related dioxygen-copper adduct [CuII(NMe2-tmpa)(Z1-O 2 )] tmpa ¼ tris(4-dimethylaminopyrid-2-ylmethyl)amine) has been spectroscopically characterized by Karlin and coworkers [61]. By starting with the

Figure 7. Schematic of two synthetic copper(II) Z1-superoxide complexes for comparison to the analogous precatalytic O2-species crystallized for PHMcc. Figure modified from [51,59,60]. Met. Ions Life Sci. 2009, 6, 295–361

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mono-carbonyl adduct [CuI(NMe2-tmpa)(CO)]1, the further reduction of 1 to form the the initially formed O2-adduct [CuII(NMe2-tmpa)(Z1-O 2 )] trans-m-1,2-peroxo dicopper(II) complex was prevented. As described in the previous section, the analogous carbonyl adduct of the parent TMPA ligand was utilized to photoinitiate transient copper(II)-superoxide formation [13].

2.2.2. Active Site Probing of the Catalytic Core of Peptidylglycine a-Hydroxylating Monooxygenase and Dopamine b-Monooxygenase through Carbon Monoxide Coordination Detailed investigations of the reaction of carbon monoxide (as a surrogate for O2) with DbM and PHMcc have been carried out by Blackburn and coworkers [54,57,62–66]. In the absence of peptide substrate, CO was found to bind to the reduced enzyme in a stoichiometry of 0.5 CO/CuI. Infrared studies and X-ray absorption spectroscopic (XAS) analysis confirmed the CO coordination occurred at the CuM site for both DbM (nCO ¼ 2089 cm1) and PHMcc (nCO ¼ 2092 cm1) (see Figure 8).

Peptidylglycine α-Hydroxylating Monooxygenase (PHMcc) N

N

N

Dopamine β-Monooxygenase (DβM) N

N 2.23 Å

N

N

CuI

CuIM

H

S

N

N

N

N

CuIM

H

N

N CO

N

N

S

N

N

N

N

N N

N tyramine N

N S

N

CuM: νCO(PHM) = 2092 cm-1 νCO(DβM) = 2086 cm-1

S

N

N

N

CO CuIM

2.25 Å

CO CuIM

H

N

N N

N

CuM: νCO(PHM) = 2092 cm-1 νCO(DβM) = 2089 cm-1 peptidyl substrate

N

CuIH CO

CuI

N

N

N

N

N N

N

2.33 Å

CO CuIM

CuIH N

N

N

N

N N

S

N

N

CO N

2.25 Å

N

CuI

N

N

N

N

N

N CuIH

CO CuIM

S

N

N N

N

CuH: νCO(PHM) = 2062 cm-1

Figure 8. Summary of the reactions of CO at the active sites of PHM and DbM and the changes occurring upon addition of peptidyl substrate to carbonylated PHMcc and tyramine to carbonylated DbM. Met. Ions Life Sci. 2009, 6, 295–361

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As is well known and documented in the prior discussions (see Section 2.1.3), infrared nCO values shift depending on the degree of p-backbonding from copper(I) to the empty p* orbitals of CO and the shift is proportional to the electron-donating ability of the ligand donor atoms. Sorrell and coworkers also have illustrated this in various synthetic inorganic systems [47,67–69]. For the pyrazole-based ligand shown in Figure 9, changes in the donor atom resulted in the following Cu(I)-CO nCO shifts: X ¼ N-amino (nCO ¼ 2082 cm1); X ¼ O-ether (nCO ¼ 2106 cm1); and X ¼ S-thioether (nCO ¼ 2123 cm1) [69]. Since the nCO value of DbM and PHMcc were higher than the values established for hemocyanin (nCO ¼ 2040–2060 cm1), CO was assumed to coordinate to CuM which has a soft sulfur donor relative to that of CuH; both CuH and hemocyanin have three imidazolyl ligands per copper. Unlike the situation seen from crystallographic data, XAS analysis conducted by Blackburn and coworkers on the oxidized and reduced forms of PHMcc indicated significant changes occur within the CuM and CuH coordination environments [53,65]. For example, upon reduction of PHMcc by ascorbate, both the CuM and CuH centers lose their solvated water ligands. Also, extended X-ray absorption fine structure (EXAFS) spectroscopy indicated that the Met314 residue was positioned 2.23 A˚ from the reduced CuM center, however was undetected (unbound) in the oxidized enzyme. In separate experiments, EXAFS data further indicated that the CuM–SMet314 distance increased from 2.23 A˚ to 2.33 A˚ upon binding CO [57] (see Figure 8). Similar changes in geometry have been observed for copper complex systems from Karlin and coworkers (Figure 9B) [11,71]. For example, the ligand-copper(I) species of the tetradentate N3S chelator ligand LN3S (LN3S ¼ 2-ethylthio-N,N-bis(pyridin-2-yl)methylethanamine) loses one coordinated

Figure 9. Copper(I)-carbonyl synthetic models with different ligand donor atoms from (A) Sorrell and coworkers [69] and (B) Karlin and coworkers [70,71]. Met. Ions Life Sci. 2009, 6, 295–361

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donor atom upon addition of CO [71]. Supporting evidence for the fourcoordinate geometry was supported by observing a very similar nCO value to that of the tridentate chelate PY1 ligand (PY1 ¼ bis(2-pyridylmethyl)amine) copper(I) carbonyl species [70]; LN3S possesses the bis(2-pyridylmethyl)amine entity. Surprisingly, the CuH site of PHMcc also coordinates carbon monoxide (Figure 8). Upon addition of peptidylglycine substrates to carbonylated PHMcc [CuM-CO..CuH], a second carbonyl stretch is observed at nCO ¼ 2062 cm1 due to CuH-CO coordination [57]. This assignment was confirmed through the use of a mutant PHMcc species (H172A) in which His172 (CuH bound) was replaced by a non-coordinating alanine residue [54,72]. Induced CuH-CO binding by substrate to the H172A mutant resulted in a 3 cm1 blue shift of the copper carbonyl [54]. Such a small CO frequency shift suggested that His172 was only weakly bound to the CuH ion of the wild-type enzyme and that CuH-CO possesses an overall three-coordinate geometry. However, the CuH-CO frequency of nCO ¼ 2062 cm1 is lower than would be expected for three-coordinate copper(I)-carbonyls, which are typically in the range 2090-2110 cm1 [54,73–75]. A change in the geometry around CuH is surprising since electron transfer proteins typically do not change their coordination number or geometry upon metal redox state changes, referred to as an ‘‘entactic’’ state. As is comparable to the CO binding studies (see just above), EXAFS of wild-type PHMcc show that one liganded CuH histidine residue (His172) is lost upon reduction, resulting in a change in the geometry from T-shaped to twocoordinate linear [53,65]. Although His172 is only weakly coordinated, the protein activity decreased to less than 1% in the H172A mutant and addition of exogenous imidazole did not restore function [54]. Blackburn and coworkers suggested that the decreased or even non-existent electron-donating ability of alanine in the H172A mutant would decrease the CuH reduction potential due to the increased stability of the linear two-coordinate geometry for copper(I) [54,74]. Another suggestion by Blackburn and coworkers for the decreased activity of the H172A mutant was that changes in the histidine coordination could alter hydrogen bond interactions critical for efficient electron transfer [54]. For example, the tyrosine residue (Tyr79) that stacks with the His172 residue could play a role in diminished activity (see Figure 6). Furthermore, replacement of the His172 residue by Tyr79 could result in the lower than expected nCO value for CuH-CO. Alternatively, decreased activity could result from a change in the His108 residue that is indirectly hydrogen bonded to peptide substrate through a glutamine residue (Gln170) as shown in the reduced X-ray structure (Figure 6). In a recent paper from Karlin and coworkers, a linear two-coordinate copper(I) complex with Nd-His ligation similar to that in CuH was reported Met. Ions Life Sci. 2009, 6, 295–361

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(see Figure 10) [74]. The synthetic complex displayed a stretching frequency of nCO ¼ 2110 cm1 that shifted to nCO ¼ 2075 cm1 upon addition of 1-methylimidazole (1-MeIm). Unlike the initial two-coordinate species, formation of a three-coordinate complex by addition of substrate enabled redox changes by allowing O2 activation. The only other synthetic threecoordinate mononuclear copper(I) carbonyl complexes with similarly low nCO values are from Sorrell and Jameson (nCO ¼ 2059–2067 cm1) [76]. In these complexes, two non-chelated imidazoles coordinate to the copper(I) species in a linear fashion and reaction with CO only occurs in the presence of excess imidazole substrate, similar to the finding for CuH. A threecoordinate binuclear copper(I) carbonyl complex of a tropocoronand macrocycle dianion (TC-5,5) characterized by Villacorta and Lippard also displayed a very low CO stretching frequency nCO ¼ 2071 cm1 (Figure 10) [77]. Other copper(I) carbonyl macrocycles such as the four-coordinate calix[6]arene-based carbonyl species from Reinaud and coworkers have much higher CO stretching frequencies of 2092 cm1 and 2102 cm1 due to their highly flexible nature [75]. In a half-apo derivative of PHMcc [CuM..( )] in which the CuH site is removed, the 2062 cm1 stretch was again present [66]. The interpretation was that copper(I) ion was transferred from the CuM site to the CuH site leading to the observed CuH-CO coordination. As alluded to above, this observation led to the proposal of a superoxide channeling mechanism that occurred between CuH and CuM that was later dismissed. A high level expression system has yet to be found for DbM and has therefore prevented any crystallographic analysis of this protein [50]. However, as mentioned, the high sequence homology and similar functions of DbM and PHM suggest similar mechanisms and coordination motifs. On the contrary, XAS and FTIR data collected by Blackburn and coworkers

Figure 10. Two-coordinate copper(I)-carbonyl adducts derived from two- or threecoordinate copper(I) complexes. The chemistry described comes from Karlin and coworkers (left) and Lippard and coworkers (right). Figures adapted from [74] and [77], respectively. Met. Ions Life Sci. 2009, 6, 295–361

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with reactions of DbM with CO and isocyanides (see below) reveals structural and chemical differences [64,78,79]. The major difference between PHM and DbM was their reactivity with CO in the presence of substrate (see Figure 8). In DbM, the CuM-CO frequency shifts by 3 cm1 (DnCO) in the presence of tyramine, potentially as a result of substrate coordination near the CuM center. However in PHM, coordination of substrate near the CuM site results in coordination of CO at the CuH site and no CuM-CO frequency shifts [63]. The same frequency shift was observed upon CuH-CO formation in the H172A mutant [54]. In addition, coordination of CO at the CuM site in DbM did not result in de-ligation of the methionine sulfur donor like in PHM [64]. Instead, displacement of an unknown fourth ligand resulted. The weakly coordinated ligand has been proposed by Blackburn and coworkers to be a nearby tyrosine residue (Tyr216 or Tyr477).

2.2.3. Coordination of Isocyanide to CuM and CuH in the Catalytic Core of Peptidylglycine a-Hydroxylating Monooxygenase and Dopamine b-Monooxygenase To further exploit the catalytic core, coordination of other copper(I) specific ligands such as isocyanides (RNC) were investigated by Blackburn and coworkers for native (PHMcc, DbM) and mutant (PHMcc) species (see Figure 11 for a schematic) [78,79]. The reduced form of PHMcc binds 2,6-dimethylphenyl isocyanide (DIMPI) as well as less bulky isopropyl

Figure 11. Reaction summary of DIMPI coordination at the active sites of PHM and DbM. No further changes are observed upon addition of a peptidyl substrate. Met. Ions Life Sci. 2009, 6, 295–361

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isocyanide (IPI) at the CuM center (nCN (DIMPI) ¼ 2138 cm1; nCN (IPI) ¼ 2174 cm1) but not at the CuH site, even in the presence of peptidyl substrate [79]. As discussed above, CO binding at the CuH site in reduced PHMcc was only observed in the presence of substrate [57]. In DbM, DIMPI coordinated to both copper centers (CuM: nCN ¼ 2129 cm1; CuH: nCN ¼ 2148 cm1), even in the absence of peptidyl substrate [78]. The contrasting reactivity of the CuH sites of PHM and DbM with isocyanides is interesting since there has been no report of CO binding to the CuH site of DbM, unlike that for PHM. Site-directed mutations of PHMcc resulted in coordination of DIMPI at the CuH site [79]. For example, replacement of the His172 residue with alanine (H172A mutant) resulted in coordination of DIMPI at both the CuM (nCN ¼ 2133 cm–1) and CuH (nCN ¼ 2147 cm1) sites. As suggested by Blackburn, the absence of a third coordinating residue to CuH could enable DIMPI binding due to redox potential changes or indirect protein structural changes [54,79]. Replacement of the CuM methionine ligand with isoleucine (M314I mutant) also resulted in coordination of DIMPI at the CuH (nCN ¼ 2149 cm1) site of PHMcc, however CuM coordination was absent [79]. Separate work by Blackburn and coworkers with the M314I mutant has shown structural perturbations that affect the overall stability of the enzyme, for example a weakening of a single CuH–histidine interaction (His107) [53,80]. Similarly, EXAFS and CO binding studies have shown that binding of substrate near CuM and/or movement of Met314 influences binding of CO to CuH (see above) [57,66]. An increase in the isocyanide to copper concentration to these enzymes results in extraction of the copper(I) ion from the protein environment and formation of a tris-isocyanide adduct [78,79]. The [(DIMPI)3-CuI]1 complex displays a single IR band at nCN ¼ 2160 cm1 and that for the corresponding [(IPI)3-CuI]1 complex is observed at nCN ¼ 2193 cm1. Even after extended dialysis of PHMcc and DbM, the IR stretch of the tris-isocyanide adduct remains, suggesting a hydrophobic interaction with the protein that prevents removal. Similar trends were observed upon dialysis of deoxygenated molluscan Hc treated with DIMPI [78]. The tridentate histidine coordination per copper site, now with coordinated DIMPI, displayed a single IR stretch at bCN ¼ 2148 cm1, identical to the DbM CuH site, that converted to 2060 cm1 with increased concentrations of DIMPI. The reaction of DIMPI with two inorganic complexes led to the formation of [CuI(MePY2)(DIMPI)]ClO4 (MePY2 ¼ bis(2-pyridylethyl)methylamine) and [Cu(1,2-Me2Im)2(DIMPI)]PF6, (1,2-Me2Im ¼ 1,2-dimethylimidazole). These were investigated by Blackburn, Karlin, and coworkers for comparison to PHMcc and DbM [78,79]. As shown in Figure 12, the structures of both have been solved by X-ray crystallography. Assignment of the DbM Met. Ions Life Sci. 2009, 6, 295–361

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Figure 12. Two-coordinate (left) and three-coordinate (right) ligand-copper(I)DIMPI synthetic models from Karlin and Blackburn. Modified from [79] and [78], respectively.

nCN values were confirmed by comparison to the four-coordinate complex [CuI(MePY2)(DIMPI)]ClO4 [78]. Consistent with the enzyme results, a clean conversion from the mono-isocyanide (nCN ¼ 2132 cm1) to the tridentate ligand displaced species [(DIMPI)3-CuI]ClO4 (nCN ¼ 2162 cm1) occurs. Preparation of authentic [(DIMPI)3-CuI]ClO4 confirmed the latter. XAS analysis of PHMcc and comparison to [CuI(1,2-Me2Im)2(DIMPI)]PF6 suggests that DIMPI is coordinated in a tilted fashion to CuM with an approximate Cu–C-N angle of 1501 in comparison to the linear model species [79]; the angle of Cu–C-N binding in PHMcc is similar to isocyanide binding in myoglobin (1501) [81].

3. HETEROBIMETALLIC COPPER-CONTAINING ENZYMES 3.1. Heme-Copper Oxidases Heme-copper oxidases (HCOs) catalyze the four-electron reduction of dioxygen to water [12,82,83]. A total of eight endogenous protons are consumed, four of which are used to produce two H2O molecules. This process is coupled to the active transport of four more protons across the lipid bilayer, creating a pH gradient across the mitochondrial membrane which is used for ATP synthesis. The active site of HCOs consists of a catalytic heterobimetallic center with the hemex3 site (x3: designated a3, b3, or o3 depending on the HCO species) positioned 4.4–5.3 A˚ from the copper (called CuB) site. The copper ion is bound by three histidine residues and the hemex3 site has a coordinated axial histidine. Higher organisms with cytochrome c oxidases (CcO) possess a Met. Ions Life Sci. 2009, 6, 295–361

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Figure 13. The structure of the cytochrome c oxidase heterobimetallic catalytic center (left). The schematic (right) summarizes the chemistry occurring following photolysis of the CcO FeII-CO species, where CO transfers to CuB, escapes and the reaction with dioxygen ensues. Modified from diagrams in [84].

copper electron acceptor site (CuA) that mediates one-electron transfer processes from cytochrome c to the low-spin hemex site (x: designated a, b, or o depending on the HCO species). Hemex then transfers electrons one at a time from CuA to the binuclear catalytic active site for O2 processing. X-ray crystal structures of various HCOs have been solved: aa3 from bovine heart (Figure 13) [85,86]; the aa3 oxidases from Paracoccus denitrificans [87] and Rhodobacter sphaeroides [88]; the ba3 oxidase from Thermus thermophilus [89]; and the bo3 oxidase from Escherichia coli (Cbo) [90]. For the remainder of this review, the catalytic active site designation has been generalized to hemea3-CuB. The last few years have seen great activity in the chemistry of HCO synthetic models [82]; only a few have been chosen for discussion herein.

3.1.1. Structural and Mechanistic Probing through Photochemical Methods Dioxygen binding to HCO active sites is too fast to follow by stopped-flow kinetic-spectroscopic methods (but see a recent advance by Wiertz et al. [91]) and thus studies to provide insight into the course of dioxygen reactivity have employed laser photoinitiated dissociation of CO from hemea3, Met. Ions Life Sci. 2009, 6, 295–361

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followed by reaction with O2 (see Figure 13). Probing O2 active-site dynamics, structure, and function, including O2 transfer between hemea3 and CuB, is also complicated by the short lifetime of intermediates. As CO can be utilized as an O2-reactivity surrogate, it has also been employed in this regard (vide infra). Through time-resolved infrared spectroscopic analysis, Woodruff and coworkers detailed the CO transfer from hemea3 to CuB, thus pointing to CuB as the entry and exit site for small molecules [92,93] (see Scheme 3). The specific mechanism involved CO rapidly switching from hemea3 to CuB upon photolysis (kFeCO/k1CuCO o1 ps), remaining transiently bound to CuB on a ms – ms timescale depending on the HCO species, before dissociating out of the active site (kCuCO). Surprisingly, CO recombination to hemea3 following dissociation from CuB has been observed only at longer timescales (k1FeCO 41 ms), even though the two metal centers are in close proximity. Until only recently was CO transfer from hemea3 to CuB fully timeresolved, through femtosecond mid-IR absorption analysis [94–96]. Previous high-resolution measurements by Vos and coworkers showed step-wise kinetics for the heme de-ligation process (kFeCO) with an initial 350 fs phase followed by a 700 fs phase [94]. The reaction dynamics were described as a heme out-of-plane motion (heme doming) following CO release. In a more recent study by Treuffet et al. on the CuB-CO ligation process (k1CuCO), the pump-probe results display an initial appearance of CuB-CO at 2064 cm1 with a delayed onset of 200 fs, followed by a 450 fs exponential rise [95]. Trajectory calculations further indicated a ‘‘ballistic’’ rather than ‘‘diffusive’’ contribution revealing an active site optimized for ligand transfer processes. Changes in the localized conformation following CO release from hemea3 (kFeCO) and/or binding to CuB (k1CuCO) (see Scheme 3) have also been suggested by Varotsis and coworkers due to the slow recombination of CO to the hemea3 site following dissociation from CuB (see Figure 14A) based on step-scan FTIR spectra collected at 298 K [98]. More specifically, CuB-CO ligation (k1CuCO) or hemea3-CO de-ligation (kFeCO) has been proposed to trigger a conformational rearrangement within the protein environment leading to a significant increase in the energy barrier for hemea3-ligand


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Figure 14. The step-scan FTIR spectrum (A) from Varotsis and coworkers and the time-resolved infrared spectrum (B) from Karlin and coworkers are representative of the same kCuCO/k1FeCO process, the thermal equilibration of Cu-CO to Fe-CO at 298 K. Spectra were modified from [97] and [84].

rebinding. As a result, the lower barrier for CO escape from the active site (kCuCO) is favored. Larsen and Miksˇ ovska´ have detailed the thermodynamics of ligand transfer in HCOs through photothermal methods such as photoacoustic calorimetry (PAC) and photothermal beam deflection (PBD) [100]. In a study utilizing PAC, the activation enthalpy (DHz) and activation volume (DVz) of ligand transfer revealed two kinetic processes (see Figure 15 for summary) [99]. The first phase is attributed to Fe-CO bond cleavage, a switch from low-spin to high-spin at the hemex3 site, and CuB–CO bond formation within 50 ns. For bovine heart CcO, the second phase is attributed to thermal release of CO from CuB in B2 ms. However, for cytochrome bo3 from E. coli, the slow phase is outside the PAC detection range and therefore at least an order of magnitude slower. An unprecedented second phase occurred within 500 ns for the HCO isolated from Rb. sphaeroides (RbCcO) that involves a volume expansion of 3.3 mL mol1 with negligible activation enthalpy (see Figure 15) [99]. This observation indicated that the process was mainly entropy-driven and was attributed to a relaxation following CO binding to CuB. The o50 ns phase in bovine heart CcO and the combined o50 ns and B500 ns events in RbCcO were thermodynamically similar, with DH B35 kcal mol1 and DV B 5 mL mol1, suggesting that conformational changes at the CuB site upon CO ligation are similar for both enzymes but occur at different timescales. However, unlike bovine CcO, the first phase for RbCcO involves no volume change and is therefore enthalpy-driven. Met. Ions Life Sci. 2009, 6, 295–361

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Figure 15. Thermodynamic profiles based on PAC studies conducted by Larsen and coworkers that depict the enthalpic (DH) changes and volume changes (DV) that occur upon coordination of CO to CuB following photoejection from Fe-CO in bovine cytochrome aa3, Rb. sphaeroides (RbCcO), cytochrome bo3 from E. coli (Cbo) [99].

Perturbations to the CuB site during the first phase in bacterial and mammalian enzymes were distinctly different according to PAC studies (Figure 15) [99]. The observed enthalpy change was roughly 12 kcal mol1 lower for Cbo in comparison to bovine heart CcO and the volume change was 12 mL mol1 lower. Previous EXAFS studies on Cbo by Blackburn and coworkers suggested that CO binding to CuB leads to an increase in the bond length between the copper(I) ion and two histidine residues [101], potentially accounting for the DHz/DVz observations obtained from PAC studies. A bio-inspired model complex, [(6L)FeII . . . CuI]1, from Karlin and coworkers, can be related to both O2 [102–104] and CO [70,84] reactivity of HCOs; 6L is a fluorinated porphyrinate with a tetradentate pyridylalkylamine appendage. Reaction of dioxygen with [(6L)FeII . . . CuI] produces a high-spin peroxo species with some similarity to that observed in one CcO X-ray structure [102–104] (see Figure 16). Coordination of CO to [(6L)FeII . . . CuI], forming [(6L)FeII(CO) . . . CuI(CO)] with nCO(Fe) ¼ 1975 cm1 and nCO(Cu) ¼ 2091 cm1, leads to a structural change in the copper moiety [70]. An additional shoulder is observed at B2070 cm1 that Met. Ions Life Sci. 2009, 6, 295–361

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Figure 16. Comparison of a synthetic heme-copper peroxo adduct (left) and a putative CcO peroxo-bridged CcO species characterized by X-ray crystallography (right). Adapted and modified from [82].

has been attributed to a second isomeric form, essentially a tridentate versus tetradentate ligand-coordination around copper(I)-CO (Figure 17). The major isomer possesses a dangling pyridine with the remaining two pyridines and central amine coordinated in a tridentate fashion to the copper(I)-CO center. The heme-bound monocarbonyl adduct [(6L)FeII(CO) . . . CuI] of the 6L binucleating ligand has been investigated by time-resolved infrared spectroscopy, displaying a photoinitiated mechanism similar to that of HCOs (see Scheme 4, Figure 14B) [84]. Photorelease of CO from the iron moiety (kFeCO) results in an immediate bleach of the coordinated carbonyl at nCO(Fe) ¼ 1975 cm1 and the appearance of a positive stretch at nCO(Cu) ¼ 2091 cm1 due to CO coordination to copper(I) upon liberation from iron(II), k1CuCO ¼ 1.5 105 s1 (298 K). Femtosecond transfer of CO to the copper metal was not observed in the 6L model system like in HCOs. Fry


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Figure 17. The two isomeric forms of a heme-copper carbonyl synthetic compound are depicted here. The major isomer (left) possesses a tridentate ligand-copper moiety. These observations exemplify how coordination number, geometry, and n(CO) values can vary. The arrow designates the point of rotation within the 6L ligand framework.

et al. suggested that the differences are likely due to competitive solvent binding or the flexible tethered portion of the 6L ligand allowing the copper(I) ion to swing away from the heme site resulting in inefficient capture of the photoreleased CO [84] (see Figure 17). The thermal equilibration of CO transfer from Cu(I) back to iron(II) is subsequently observed, kCuCO/1FeCO ¼ 1600 s1 (298 K) [84]. The rate of this second process (kCuCO/1FeCO) is within range of values measured for CO transfer from CuB to hemex3 in HCOs: aa3 ¼ 1030 s1; caa3 ¼ 50 s1; and ba3 ¼ 29 s1 [84,97,105–111]. Additionally, the enthalpic barrier, DHz ¼ 43.9 kJ mol1 (kCuCO/1FeCO), is remarkably similar to that obtained upon CO photolysis from the analogous mononuclear species [CuI(tmpa)(CO)]1, DHz ¼ 43.6 kJ mol1 [11]; suggesting that CO dissociation (kCuCO) from copper(I) regulates the binding of small molecules to the heme in the model system as well as in natural systems.

3.1.2. Proton Translocation Pathways Various proton pumping pathways (K, D, H, Q) have been proposed based on reported X-ray crystal structures and mutagenesis studies on CcOs [83]. The CuB site has been believed to play a role in the proton translocation process due to structural changes observed in nearby amino acid side chains upon transiently binding CO. One of the three histidine residues coordinated to CuB is covalently linked to a tyrosine residue, forming a His-Tyr crosslink (see Figure 13). Among various proposals that have appeared, one is that the Met. Ions Life Sci. 2009, 6, 295–361

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His-Tyr crosslink provides an initial site for electron/proton transfer during the O2 reduction process. IR spectroscopy has been the major tool employed to investigate ligation-coupled protonation/deprotonation events of protein carboxyl groups [106,112–119]. Varotsis and coworkers have utilized time-resolved step-scan FTIR and density functional theory (DFT) to investigate changes in the CuB-CO environment under variable pH/pD, ranging from approximately 5.5–10 [112,113]. In T. thermophilus HCOs, the nCO frequency at 2053 cm1 was invariant revealing the rigidity of the CuB site [106,113]. However, in Pseudomonas stutzeri, the nCO ¼ 2065 cm–1 value shifted by 1–5 cm1 upon changes in the pH/pD [112]. Histidine deprotonation (DnCO ¼ 20–43 cm1) or de-ligation (DnCO ¼ 38–121 cm1) events at the CuB active site would result in significant nCO shifts. Therefore, the observed small frequency shifts were assigned to conformational changes in the vicinity of CuB and the function of the His-Tyr crosslink was assigned as a means of fixing the distance of the CuB site from hemea3 [114], as suggested originally by Das et al. [237]. In a separate study, resonance Raman (rR) spectroscopy was used to detect a photostable five-coordinate hemea3-CO adduct of T. thermophilus [115]. De-ligation of the axial Ne-His384 residue from hemea3-CO disrupts the hydrogen bonding between Nd-His384 and the carbonyl of Gly359, which would in turn affect the Q-proton pathway. Yoshikawa and coworkers monitored changes upon CO and CN– coordination to the hemea3-CuB site by FTIR in order to determine the redox active metal responsible for changes in the Asp51 confirmation (H-pathway) [116]. The difference FTIR spectra of fully oxidized versus fully reduced enzyme gave a peak at 1738 and 1585 cm–1 attributed to Asp51. Identical spectra are observed upon binding CO to the reduced species (FeII-hemex3, CuIB) and CN– to the oxidized species (FeIII-hemex3, CuII B ). Since CO coordinates to both the CuB and hemea3 site in the reduced enzyme (vide infra), and CN– binds to the hemea3 site of the oxidized enzyme only, neither are believed to control the protonation state of Asp51 in the H-pathway, instead the CuA and/or hemea sites were the proposed candidates. However, mechanistic variations exist amongst different CcO species and/or pathways. For example, upon formation of CuB-CO following photolysis from hemea3, the hydrogen bonding of Glu278 in P. denitrificans [117], or Glu286 in E. coli cytochrome bo3 [118,119] and Rb. sphaeroides [120] is altered, which would in turn have an effect on the proton gating D-pathway.

3.1.3. Mixed-Valent Heme-Copper Oxidases and Electron Transfer Photoinitiated ejection of CO from hemex3 to CuB has also been used to examine the back electron transfer from hemex3 to hemex. The experiments Met. Ions Life Sci. 2009, 6, 295–361

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discussed thus far have involved fully reduced HCOs (FeII-hemex3, CuIB, FeII-hemex, CuICuIA). Photolysis of CO from FeII-hemex3 and transfer to CuIB inherently lowers the redox potential of the hemex3 site. Therefore, in mixed-valent HCOs (FeII-hemex3, CuIB FeIII-hemex, CuIICuIA), photoejection of CO induces rapid electron transfer from ferrous hemex3 site to ferric hemex. In a study by Vos and coworkers [121,122] on mammalian CcO, transfer of CO to CuB is followed by rapid electron redistribution between hemex3 and hemex (B1.2 ns) (see Figure 18). Further electron transfer from hemex3 and hemex occurs in B3 ms which is on the same timescale as the thermal release of CO from CuB in the fully reduced enzyme, suggesting a coupled electron transfer/ligand release process (see Figure 18). These types of experiments further illustrate the broad extent of insights into CcO structure and function obtainable from heme-CO photolysis chemistry. At any step, in order to move electrons to the binuclear active site, the reduction potential of hemex3 must remain higher than hemex (see Figure 19) [123]. In the final step of the catalytic cycle for O2 processing by CcO,

Figure 18. Summary of the electron redistribution process that occurs between hemea and hemea3 upon the photoinitiated transfer of CO to CuB, and the potentially coupled CO release from CuB and electron transfer process. Diagrams adapted from [96] and [100]. Met. Ions Life Sci. 2009, 6, 295–361

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Figure 19. Scheme A summarizes a proposed redox-coupled proton transfer mechanism at the HCO active site, modified from [123]. Scheme B exemplifies the insensitivity of FeIII to CuI without a bridging species such as cyanide.

i.e., following full reduction of O2 to produce two H2O molecules, the binuclear center is in the fully oxidized state (O state). After a first electron transfer from hemex to form FeIII-hemex3/CuIB (E state), the reduction potential of ferric hemex3 is significantly lowered, making the second electron transfer to form the fully reduced FeII-hemex3/CuIB state (R state) difficult (see Figure 19A). Through spectroelectrochemical studies conducted on an engineered heme-copper center of myoglobin, Lu and coworkers showed that redox changes at the copper site had little effect on the heme unless a bridging molecule such as CN– was present, see Figure 19B [123]. Thus, it is likely that compensation by a positive charge or proton would be necessary to facilitate the next electron transfer in the enzyme, supporting the proposal that redox-coupled proton transfer regulates the reduction potential of hemex3 during catalysis (see Figure 19A). FTIR studies on photolysis of the carbonyl adduct of mixed-valent species of bovine CcO have linked the protonation state of Glu242 (pathway D) to the redox state of hemex3/hemex suggesting a correlation with proton transfer events [124,125]. In a more recent EPR study by White et al. on a mixed-valent carbonyl adduct of mutant cytochrome bo3 from E. coli, arginine residues positioned at the cytosolic entrance to the D- and K-proton channels, Arg134 and Arg309 respectively, were replaced by nitroxide spin labeled cysteine residues in order to monitor for conformational changes [126]. Arg134 is adjacent to the aspartic acid (D) residue Asp135 that is essential for the D-pathway, while Arg309 is positioned near the leucine (K) residue Leu362 that is essential for the K-pathway (see Figure 20). However, there was no evidence for protein motions at the entrances to the channels Met. Ions Life Sci. 2009, 6, 295–361

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Figure 20. X-ray crystal structure of Cbo subunit I with specific residues marked along the D- and K-proton pathways (see text for relevance). Adapted from [126].

supporting that redox gating of protons occurs deep within the protein matrix; for example at Glu286 as was discussed in the previous section.

3.1.4. Interaction of Cyanide Cyanide inhibits cellular respiration and ATP production by binding to the hemea3-CuB binuclear center [127,128]. Such inhibition produces oxidative stress due to the generation of excess reactive oxygen species contributing to cellular dysfunction and eventual death if untreated. Although molecular oxygen can bind to the fully reduced (FeII-hemex3-CuIB) enzyme only, CN can react with the fully reduced, fully oxidized (FeIII-hemex3-CuII B ), and ) forms [129,130]. partially reduced (FeIII-hemex3-CuIB or FeII-hemex3-CuII B In fact, cyanide most efficiently traps the binuclear center of CcO in the partially reduced state. In bovine heart cytochrome aa3, the electron is primarily on the copper atom in a FeIII-(CN)-CuIB trapped state [131]. On the other hand, in T. thermophilus cytochrome ba3, the electron is primarily on the iron atom in a FeII-(CN)-CuII B trapped state. The different electron Met. Ions Life Sci. 2009, 6, 295–361

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Figure 21. Various heme-copper bridged assemblies that utilize the same heme moiety with different copper(II/I) precursors resulting in different Fe-CN-Cu or FeNC-Cu conformations. Adapted from [133].

distributions are believed to correlate with their proximal histidine-iron distance of 1.9 A˚ for aa3 and 3.3 A˚ (unbound) for ba3 [85–87,132]. However, Nicholls and Soulimane suggested that key events related to redox changes occur at the CuB site based on electronic changes upon binding cyanide [131]; similar work was previously conducted by Oertling et al. [132]. The large differences in equilibrium constants (KCO) for CO binding to CuB as well as the CO dissociation rates (kCuCO) in the aa3 (KCO ¼ 4 106 M1; kCuCO ¼ 0.02 s1) and ba3 (KCO ¼ 1 105 M1; kCuCO ¼ 0.8 s1) enzymes also support this proposal [111]. The mode of CN– binding in HCOs, whether terminal or bridged, has yet to be completely confirmed. Lim and Holm have carried out extensive studies on molecular heme-cyanide-copper bridged assemblies [133]. A general trend (Figure 21) was established for the linkage isomers Fe-CN-Cu and Fe-NC-Cu, based on the linearity of the species, the C-N stretching frequency, and the spin state. If the angle involving iron is within or close to the range 175–1801 but the angle involving copper is not, then the bridge is likely Fe-CN-Cu; otherwise, if the converse is true concerning angles, then the bridge is likely Fe-NC-Cu. The IR criteria for the species fall within the Met. Ions Life Sci. 2009, 6, 295–361

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Figure 22. Terminal CcO-cyanide complexes reported by Tsubaki and Yoshikawa with different coordination modes. Adapted from [135].

ranges listed in Figure 21. Three X-ray crystal structures have been solved for Fe-NC-Cu assemblies, a schematic of these species is shown in Figure 21 [82,133,134]. A Fe-CN-Cu bridged structure has been the most discussed for the natural systems, whereas the Fe-NC-Cu has yet to be proposed. For synthetic model systems, the heme coordination in the Fe-CN-Cu isomers are six coordinate as shown in Figure 21, as opposed to the five-coordinate structures exhibited by the Fe-NC-Cu isomers [133]. As mentioned above, the difference in reactivity of cytochrome aa3 and ba3 was attributed to the heme proximal histidine bond distance [85–87,132]. Therefore, it seems reasonable to consider the existence of a Fe-NC-Cu bridge in some enzyme forms. Terminal cyano-copper protein complexes have also been observed. For example, through FTIR spectroscopy, Tsubaki and Yoshikawa examined cyanide binding at the CuB site in aa3 CcO from bovine heart [135]. Under high concentrations of CN, two cyanide stretching frequencies attributed to CuB-(CN)n were observed. The more prominent nCN value at nCN(Cu) ¼ 2093 cm1 was attributed to a monocyano-copper species (CuB-CN). The conformation was assigned as a terminally bound CN pointed away from hemea3 (Figure 22). Competition experiments using CO further supported cyanide coordination at the copper site. Futhermore, the nCN value for CuB-CN remained unchanged with FeIII/II redox changes. A dicyano-copper complex CuB-(CN)2 was also observed that exhibited a second nCN(Cu) ¼ 2037 cm1 stretching frequency. The conformation was assigned as an orientation pointed toward the hemea3 site (Figure 22). The second nCN value was not observed at low cyanide concentrations nor in the presence of CO.

3.1.5. Nitric Oxide Reduction Capabilities of Cytochrome c Oxidase Nitric oxide (NO) is a cell signaling molecule responsible for mammalian immune response that can also inhibit the hemea3-CuB binuclear center Met. Ions Life Sci. 2009, 6, 295–361

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[136]. At high physiological concentrations, NO can react with radical oxygen species such as superoxide (O 2 ) to produce toxins such as peroxynitrite (OONO) [137]. Alternatively, if superoxide dismutases (SODs) (see below) are impaired or saturated due to inhibition of CcO by NO then OONO can also be formed [138,139]. Therefore, the capability of mammalian CcOs to metabolize NO to N2O like analogous bacterial NO reductases (NOR) is of physiological importance. Differences in NO reduction capabilities amongst HCOs is thought to be related to NO binding affinity of CuB; this supposition comes from the observations that the coordination affinity of CO is much higher in some bacterial HCOs (KCO ¼ 103–104 M1) in comparison to mesophilic oxidases (KCO¼101–102 M–1) [111,137,140]. It has been suggested that the higher affinity of bacterial HCOs for binding CO and presumably O2 (or NO) is a characteristic that has evolved as a result of the typical environmental conditions, i.e. low gas solubility at high temperature [111]. In recent work by Leavesley et al., the intracellular versus extracellular concentration of NO in mammalian CcO was shown to have antagonistic effects on cyanide-mediated toxicity [128]. At low endogenous levels, NO II and CN– additively inhibit CcO through CuII B -NO and Fe -CN coordination. At high NO concentrations, cyanide is displaced from the binuclear active site, and then NO coordinates to both metal centers. As a result, NO is reduced to nitrite and cyanide toxicity decreases, as initially demonstrated by Pearce et al. [128,141]. As deduced from CO binding studies mentioned above, the CuB site is believed to play a role in altering the active site accessibility. For example, by acting as the doorway for substrates both in and out of the active site or as a consequence of effects upon nearby amino acid side chains. In a biomimetic model study by Collman and coworkers, the presence of a copper ion near a heme decreased the susceptibility of the CcO model to inhibition by CN and CO during O2 reduction chemistry [142].

3.2. Cu-Zn Superoxide Dismutase Copper-zinc superoxide dismutase (CuZnSOD) is responsible for relieving oxidative stress by catalyzing the disproportionation of superoxide into less toxic products, dioxygen and hydrogen peroxide [143–145] (Scheme 5). Mutations in the protein are linked to the neurodegenerative disease amyotrophic lateral sclerosis (ALS or Lou Gehrigs disease). As such, this enzyme continues to attract great research interest and activity. The active site of the oxidized form of bovine CuZnSOD consists of a fivecoordinate copper(II) ion bound by an axial water molecule and four histidyl residues (His44, His46, His118, and His61) that form a distorted square Met. Ions Life Sci. 2009, 6, 295–361

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Scheme 5.

plane [146–148]. The zinc ion is coordinated by three histidines (His69, His78, and His61) and one aspartate residue (Asp81). The two metal ions are bridged by the deprotonated His61 (imidazolate) residue (Figure 23). The imidazolate bridge is believed to have a key role in facilitating the near diffusion controlled scavenging of superoxide (109–1010 M1s1) by stabilizing the redox active copper(I/II) center [149,150]. Second-shell ligands are also believed to play a role through electrostatic trafficking [151–153]. A basic enzymatic mechanism for CuZnSOD involves two sequential steps that are first order in superoxide [145,154]. First, the copper(II) ion is directly reduced by O 2 through inner sphere electron transfer, releasing dioxygen. The second superoxide is proposed to hydrogen bond to a nearby arginine (Arg141) and indirectly reoxidize the now CuI ion through outer sphere electron transfer, generating hydrogen peroxide. Structural and spectroscopic analysis have suggested that after substrate release in the reduced CuI form of the enzyme that the imidazolate becomes protonated and moves away from the copper center by o1 A˚ [146–148]. In a number of studies, cyanide ion has been used to probe aspects of solution coordination at the CuZnSOD active-site [155–159]. Early work by Haffner and Coleman provided evidence for cyanide binding through the carbon atom to the copper(II) site of CuZnSOD through electron spin resonance spectroscopy using a series of isotopically labeled cyanides (i.e., 12 14 12 15 C N , C N , and 13C14N) [159]. More recent studies have utilized

Figure 23. X-ray structures of the active site of bovine CuZnSOD in the oxidized (left) and reduced (right) forms; the imidazolate bridge is broken upon reduction. Modified from [145]. Met. Ions Life Sci. 2009, 6, 295–361

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ultraviolet resonance Raman spectroscopy (UVrR) to monitor changes in metal-bound histidine ligation upon surrogate (CN) substrate binding [155–159]. Metal-bound imidazolate exhibits a characteristic 1282/ 1292 cm1 doublet band attributed to the bridged conformation, and a single band at 1564 cm1[155]. Upon coordination of cyanide, evidence for copper(II) reduction to copper(I) and copper(I)-His61 deligation is observed based on changes in the UVrR spectra. The doublet characteristic of the imidazolate bridging conformation collapses to a single band at 1287 cm1 [156–158]. In addition, there is a downward shift of the 1564 cm1 band associated with metal-bound His61. Spiro and coworkers have suggested that the spectral changes are due to the axial bound water molecule being replaced by cyanide. The tetragonal geometry around copper rotates resulting in a reorientation of the His61 imidazolate ring, placing cyanide in an equatorial ligand position [158]. Such work supports that the imidazolate bridge facilitates copper redox interconversions by modulating the ligand field around the CuII/I ion.

3.3. Molybdenum-Copper Carbon Monoxide Dehydrogenase The carbon monoxide dehydrogenase isolated from the eubacterium Oligotropha carboxidovorans was recently recognized as the first molybdenum and copper containing metalloenzyme [160]. The function of the Mo/Cu-dependent protein (MoCuCODH) is to maintain atmospheric carbon monoxide levels while supplying energy to the organism through the conversion of CO to CO2. X-ray structural characterization of MoCuCODH has been reported, however the detailed mechanism is still under debate [160–163]. X-ray crystal structures have been solved for the copper-depleted inactive form, the oxidized, and the reduced states of MoCuCODH [160]. The dinuclear protein consists of a two-coordinate copper center bridged by sulfide to a five-coordinate molybdenum center. The copper(I) ion is also bound by a terminal cysteinate. In both the oxidized and reduced state, the MoVI/IV ion is chelated by two sulfur atoms of a metallopterin (mtp) ligand (as found in all mononuclear molybdenum enzymes), an axial oxo group and an equatorial hydroxyl group as shown in Figure 24. The ‘‘active’’ coordination environment and possible metal oxidation states were confirmed by X-ray absorption spectroscopy [161]. The position of the Mo and Cu K-edge revealed that molybdenum is the site of redox activity; the copper(I) center is redox inactive. The active oxidant form of MoCu CODH was determined as a [CuI(m-Scys)MoVI(¼O)2] cluster that converts to a [CuI(m-Scys)MoIV(¼O)-OH(2)] cluster upon reduction and CO Met. Ions Life Sci. 2009, 6, 295–361

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Figure 24. [164].

333

X-ray crystal structure of the active site of

MoCu

CODH. Adapted from

oxidation. Copper(I) is known to reversibly form copper(I)-carbonyls and has therefore been postulated as the substrate binding site. Dobbek et al. discussed a reaction pathway for CO oxidation in MoCu CODH based on the similar reactivities of CO and isocyanide in metallochemistry and the observation and isolation of a tert-butyl isocyanide (t-BuNC) inhibited enzyme form [160]. The proposed mechanism involved an initial fully oxidized species, [CuI(m-Scys)MoVI(¼O)-OH] followed by an ‘‘open state’’ intermediate formed by insertion of CO between copper(I) and the sulfido, hydroxy ligands of molybdenum(VI) to produce a thiocarbamate bridge between CuI and MoIV (see Figure 25). Upon release of CO2 and substitution by H2O, the Cu-m-Scys bond is reformed, producing [CuI(m-Scys)MoIV(¼O)-OH]. The thiocarbamate bridge ‘‘open state’’ intermediate (see Figure 25) was termed based on the increase in the molybdenum copper distance from 3.74 to 5.07 A˚ following the insertion chemistry [160]. Since this structure-based proposal, two revised mechanisms have been reported based on DFT calculations that take into consideration the energetics of the various steps [162,163]. In both of these computational

Figure 25. Proposed ‘‘open state’’ intermediate based on X-ray structure of MoCu CODH with bound t-BuNC. Modified from [162]. Met. Ions Life Sci. 2009, 6, 295–361

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Figure 26. MoCuCODH mechanism proposed by Kirk and coworkers, based on their observed electronic ‘communication’ between Mo and Cu, and standard metalCO binding and insertion chemistry [165].

studies, the bis-oxo form of the active center [CuI(m-Scys)MoVI(¼O)2] is considered the catalytic site, even though the direct pathways for CO oxidation differ. From a detailed spectroscopic study, Kirk, Young, and coworkers showed that electrons released during the reaction pathway likely transfer via the sulfido ligand to yield the reduced Mo(IV) state (Figure 26). Extensive electronic delocalization exists in a synthetic MoV(m-S)CuI model compound, as indicated by the highly rhombic EPR spectra with strong 63,65Cu hyperfine coupling constants (A B 60 104 cm1) in frozen glass [164]. Both the native enzyme and the protein model undergo cyanolysis, i.e., CN reactions which break the Mo-S-Cu bond. While not discussed in this report, the probable copper product is the known [CuI(CN)3]2 ion [166]. Based on these findings, along with the known ability of copper(I) to coordinate CO and the fact that metal-carbonyls undergo nucleophilic attack at the carbon atom, Kirk and coworkers propose a relatively simplified reaction mechanism for CO oxidation outlined in Figure 26 [164,165].

4. NON-BLUE COPPER OXIDASES 4.1. Copper Amine Oxidase Copper amine oxidases (CAOs) are ubiquitous enzymes found in both prokaryotes and eukaryotes that oxidize primary amines to their corresponding aldehyde (see Figure 27). The copper(II) active site structure of CAO from Arthrobacter globiformis (AGAO) is five-coordinate, consisting of a water molecule and three conserved histidine residues bound in the equatorial position and one axially bound water [167]. According to X-ray absorption studies, the coordination number of the CAO copper ion decreases from five (3 Ne-His, 2 H2O) to three (includes at least 2 Ne-His) upon reduction [168]. An organic cofactor (TPQ ¼ 2,4,5-trihydroxylphenylalanine quinone) that is Met. Ions Life Sci. 2009, 6, 295–361

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Figure 27. X-ray crystal structure (left) of the CAO active site, including the TPQ cofactor; modified from [3]. The reaction schemes (right) include the biogenesis of TPQ (top) and the reductive and oxidative half-reactions for amine oxidation (bottom).

critical for enzyme function is positioned close to the copper(II) ion [169]. The TPQ cofactor is formed by a copper catalyzed, O2-dependent, posttranslational modification of a tyrosine residue [170]. The catalytic cycle of CAO follows a ping pong mechanism with reductive and oxidative reactions as shown in a condensed form in Figure 28. The amine is oxidized to aldehyde in the first phase, generating the two-electron reduced form of the cofactor (TPQred). In the second step, the TPQred cofactor is reoxidized (TPQox) and molecular oxygen is reduced to hydrogen peroxide. The pathway of the first electron transfer in the latter step is still intensely investigated; two alternative pathways are shown in Figure 28. Pathway 2, which involves direct transfer of an electron from the TPQred cofactor to molecular oxygen, had been increasingly supported [171,172]. However, another possible mechanism, labeled pathway 1, involves the initial transfer of an electron from the TPQred cofactor to the copper(II) center, generating copper(I) and a radical semiquinone cofactor species (TPQsq); this is followed by rapid Cu(I)–O2 activation, i.e., dioxygen binding to copper(I) and reduction [173,238]. An important difference of the two aforementioned mechanisms is the position of the TPQred/CuII to TPQsq/CuI equilibrium. In support of pathway 2 (that first stated), recent data has indicated that changes in the level of TPQsq/CuI, which varies from almost 0% to 40% depending on the enzyme source, is not enough to alter the mechanism of O2 reduction [172]. The equilibrium position as a function of pH was measured by addition of cyanide to the anaerobic mixture of TPQred/CuII and TPQsq/CuI which pushes the equilibrium to TPQsq/CuI. TPQsq has distinct visible features at 360, 435, and 465 nm, whereas TPQred does not absorb above 350 nm. Therefore, the relative position of the equilibrium could be Met. Ions Life Sci. 2009, 6, 295–361

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Figure 28. Two proposed oxidative half reactions labeled pathway 1 and 2. Adapted from [172].

observed by monitoring changes in the 465 nm absorption, before and after reduction. In a separate study, the role of cyanide in the inhibition of CAO reactivity was investigated [173]. Addition of cyanide resulted in cyanohydrin derivitization of the quinone cofactor. This did not significantly affect the mechanism for inhibition towards substrate amine. Instead, complexation of cyanide and copper(I) played the major role in inhibition. The presumed explanation was CN/O2 competition that prevented reoxidation of TPQred, supporting pathway 1; a potential equilibrium perturbation to support pathway 2 was mentioned. Carbon monoxide was used as a probe to gain further insight about the copper(I) CAO structural environment. Hirota et al. generated copper(I) carbonyl adducts of AGAO with TPQ in different redox states [174]. Changes in the chemical and redox state of TPQ resulted in significant variations of the copper(I)-carbonyl nCO values. Two CO-binding modes were observed in each system as follows: 2064 cm1 and 2083 cm1 for Met. Ions Life Sci. 2009, 6, 295–361

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TPQox/CuI-CO; 2063 cm1 and 2079 cm1 for TPQsq/CuI-CO; 2061 cm1 and 2077 cm1 for TPQred/CuI-CO. The lower nCO frequencies at 2061– 2064 cm1 emerged during the beginning of the substrate reduction process and were believed to represent CO substitution for an equatorial water. The higher nCO frequencies at 2077–2083 cm1 were thought to arise from copper coordination changing from three to two imidazoles. The results support that the copper(I) ligand coordination is modulated by the chemical and redox states of the TPQ cofactor. A change in the copper(I) coordination number was originally suggested by Dooley et al., based on XAS studies [168].

4.2. Galactose Oxidase Galactose oxidase (GO) catalyzes the oxidation of primary alcohols into their corresponding aldehydes with concomitant reduction of dioxygen into hydrogen peroxide [175]. The active site of GO is comprised of a copper(II) ion coordinated to imidazole (His) and phenol(ate) (Tyr) residues, and additionally to the phenolate group of a post-translationally modified, crosslinked cysteine-tyrosine radical cofactor [3]. The oxidative coupling mechanism responsible for formation of the Cys-Tyr crosslink is not well understood. Another interesting structural feature is the stacking of a tryptophan residue with the Cys-Tyr cofactor, which plays a multifunctional role in enzyme catalysis from substrate binding to tyrosyl radical redox potential and stability [176,177]. GOs are still actively studied [175]; however, detailed, now well established, enzyme reaction chemistry will not be discussed herein because little ‘Cu-C’ literature exists. Cyanide was used to aid in probing exogenous ligand binding to the active site of GO via nuclear magnetic relaxation dispersion (NMRD) measurements [178]. Solvent water exchange, inhibitor (CN–) and substrate binding were all found to occur at the equatorial position.

5. BLUE, GREEN, AND PURPLE COPPER PROTEINS 5.1. Blue Electron Transfer Proteins ‘Cu-C’ interactions are rare in blue copper proteins, which function as electron-transfer mediators in important biochemical processes such as photosynthesis and metabolism [179,180]. Classified as type 1 (T1) proteins, they are characterized by an intense charge-transfer transition centered around 600 nm (e ¼ 2,000–10,000 M1cm1), high CuII/I redox potentials (+180 to +800 mV), and display unusually small copper parallel hyperfine Met. Ions Life Sci. 2009, 6, 295–361

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splittings (AJ ¼ 43–95 104 cm1). The highly conserved ‘‘blue’’ active site consists of a copper(II) center coordinated by one cysteine and two histidine residues in a trigonal array. The intense 600 nm absorption is believed to arise from excellent CuII-Scys orbital overlap (Scys p - Cu dx2y2) reflected by the short 2.1–2.3 A˚ bond distance. A charge transfer transition at higher energy, typically around 450 nm, is also due to the CuII-Scys orbital overlap (Scys (pseudo) s - Cu dx2y2). This strong covalency removes unpaired electron density on the copper nucleus to the Scys-ligand leading to the small hyperfine coupling AJ value. In small proteins with a single ‘‘blue’’ copper site such as plastocyanin (Pc) and azurin (Az), a weakly interacting methionine residue is located in the axial position at 2.6–3.1 A˚ [181]. Az additionally coordinates to the backbone oxygen of a glycine residue, also positioned axial and at approximately the same distance (B3.1 A˚) from the copper center. The long, potentially non-bonding SMet distance in Pc and Az is believed to contribute to the reduction potential of the copper center, as suggested by observed differences when axial ligands are altered via site-directed mutagenesis experiments. The ability of the axial methionine to prevent unwanted reactions from occurring with exogenous ligands has been examined through optical and electron paramagnetic resonance spectroscopy. Small molecules (cyanide) can in fact coordinate to the copper center in Az from Pseudomonas aeruginosa when the axial Met ligand is replaced with non-coordinating residues such as glycine or alanine [182]. With now stronger axial interactions due to coordination of cyanide (instead of the typical Met ligand), the two visible optical absorbances shift in parallel to higher energy and the intensity of the 430 nm absorption increases at the expense of the 600 nm absorption; this demonstrates their identical origin. The proposed explanation is that the stronger axial coordination results in displacement of the copper ion from the Cys-His-His plane leading to decreased overlap between the Scys p and Cu dx2y2 orbitals and an increase of the overlap between the Scys (pseudo) s and Cu dx2y2 orbitals. EPR data analysis showed that the gz value decreases with strong axial ligation.

5.2. Multi-Copper Oxidases Naturally occurring blue multi-copper oxidases (MCOs) are multi-domain enzymes that have a minimum of four copper(II) centers, consisting of a T1, type 2 (T2) center and a dinuclear type 3 (T3) site [39,183]. The T1 copper center exhibits features very similar to the blue electron transfer proteins, see above. The T2 copper center does not possess distinctive absorbance features and exhibits ‘normal’ EPR parameters (AJ ¼ 158–201 104 cm1). The three-coordinate geometry of T2 sites is unusual because it consists of Met. Ions Life Sci. 2009, 6, 295–361

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only two histidines and a hydroxide or water ligand. The dinuclear T3 site is EPR-silent as each copper(II) ion binds three histidine ligands and a bridging hydroxo or oxo group. Ascorbate oxidase (AO; 3 domains), laccase (Lc; 3 domains), and human ceruloplasmin (Cp; 6 domains) are blue multi-domain copper oxidases that catalyze the four-electron reduction of dioxygen to water (see Scheme 6). Electrons are supplied by substrates such as ascorbate, phenols, and aromatic amines; iron(II) is a substrate for Cp. The Cu-SMet axial ligand typical for blue T1 sites such as in AO and Lc is replaced by a leucine residue. The greater hydrophobicity about this T1 copper environment leads to the higher redox potentials for Lc (+785 mV versus NHE) and the T1 site in the second domain of Cp (B+1,000 mV versus NHE). The role of the MCO blue T1 centers is to harvest electrons from substrate oxidation and transfer these to the trinuclear T2/T3 center. The T2 and T3 sites form a trinuclear cluster and function as the site of dioxygen binding and reduction to water. See Figure 29 for a relevant MCO structure of ascorbate oxidase. To probe MCO active site ligand coordination, the effect of enzyme inhibitors (cyanide) on the kinetics of T1 reduction and intramolecular electron transfer from T1 were measured through laser flash photolysis studies for AO from zucchini squash [184]. The reduction of the T1 site by lumiflavin semiquinone was measured as 2.7 107 M1 s1 and decreased by a factor of two in the presence of cyanide. The dissociation constant for cyanide (0.2–0.4 mM) was similar to other reported inhibition constants for AO (0.13–0.46 mM). The same experiment was conducted for an AO mutant in which the copper ion from the T2 site was removed. The rate constant for dissociation was the same for both the native and copper depleted form, suggesting cyanide binds to the binuclear T3 site. Additionally, the rate constant for intramolecular electron transfer was the same suggesting that


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Figure 29. Structure of AO exemplifying the multicopper active site, including the T1 site and the T2/T3 triad. Coordinates (1AOZ) were taken from the Protein Data Bank (Brookhaven) and displayed using the program Rasmol.

the T3 center is the immediate acceptor of the first electron from the T1 copper. There is a great deal now known concerning the O2 reduction chemical mechanism in MCOs, in large part due to the research of Solomon and coworkers [39,180]. As mentioned, Cp catalyzes the oxidation of FeII to FeIII and is believed to play a role in iron metabolism. Cp also carries copper ion in plasma and is believed to play a part in copper transport/regulation. In comparison to AO and Lc, Cp is unique because it contains six copper ions rather than a multiple of four. More specifically, there are three T1 sites per T2/T3 cluster which has been proposed to have a functional role. Treatment of Cp with cyanide has shown that there is a preferential loss of T1 copper rather than binding at the T2/T3 dioxygen reduction sites [185,186]. The observation that the T1 site is labile suggests that the enzyme can reversibly release some of its intrinsic copper without completely sacrificing oxidase activity. Lc has been proposed as a potential candidate for the detoxification of environmental pollutants due to its ability to degrade lignin in kraft pulps, an important step in the mineralization of carbon in nature [187]. Lc has also been shown to catalyze the oxidation of triclosan, an antibacterial agent that is widely used in the commercial industry [188]. However, cyanide as a potential contaminant in wastewater has been shown to inhibit the Lc lignin activity [189,190].

5.3. Copper Enzymes in Denitrification 5.3.1. Nitrite Reductase Copper nitrite reductases (NIRs) have a functional unit consisting of a T1 and T2 site and they are classified into two groups based on the T1 Met. Ions Life Sci. 2009, 6, 295–361

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spectroscopic properties, which are slightly modified from T1 sites from electron transfer proteins (see above). They all possess 2 His, 1 Cys, and 1 Met ligand, but the Blue reductases have an axially distorted tetrahedral arrangement and the Green reductases have an axially flattened rhombically altered metal center. The catalytic center which is accessible to neutral or anionic small molecules, including the substrate nitrite, is coordinated by three His residues and a water molecule in a tetrahedral geometry. The T1 center passes electrons to the nearby T2 site to effect the one-electron 1 reduction NO 2 +e +2H - NO+H2O. As would be expected for an active site that allows small molecules to enter or exit and where both reduced and oxidized Cu(I) and Cu(II) can reside, both CO and CN are NIR inhibitors. As such, Mauk, Murphy, and coworkers used CO to probe the active site of NIR from Alcaligenes [191]. Carbon monoxide binds to the reduced enzyme, and in the wild-type NIR it exhibits a single CO stretch at 2050 cm1. However, in an Asp98Asn mutant enzyme, two bands were observed, at 2060 and 2041 cm1. The latter shifts down by 2 cm1 in D2O, suggesting it is subject to H-bonding effects. They further deduce that in the native enzyme, the copper-carbonyl moiety is not subject to such influences. Overall, the authors concluded that Asp98 is key in organizing the orientation and dynamics of ligand binding to the active site copper. Based on known NIR X-ray structures, they constructed a model of CO bound to the active site (Figure 30). As implied by the discussions concerning copper-carbonyl complexes and stretching frequencies, they are certainly highly subject to coordination environmental factors such as the number of ligand groups (e.g., 3 N versus 4 N ligands), the coordination geometry, along with the identity (e.g., S versus N) and donor strength of the ligands. We note for example a series of

Figure 30. Model constructed of the NIR T2 active-site with Cu(I)-CO moiety (left, adapted from [191]), and diagram of the LR ligands and n(CO) values of derived Cu(I)-CO complexes (right). Met. Ions Life Sci. 2009, 6, 295–361

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Figure 31. A side-on bound copper-nitrosyl observed in a NIR protein structure. Adapted from [192].

ligand-copper(I) complexes LR (Figure 30), which vary systematically in donor strength as a consequence of 4-substituents on pyridyl groups. The CO stretching frequency varies considerably just due to this donor effect; the stronger donor(s) increase back-bonding to the CO p* orbital, lowering the n(CO) value. It is worthwhile to note that protein active site n(CO) values are found at generally lower frequencies (2043–2063 cm1) than for synthetic complexes with neutral ligands (2080–2102 cm–1) [75,192]. This suggests that copper proteins with several histidine imidazole ligands are stronger donors than for example aliphatic amines or pyridines. Copper complexes with nonchelating imidazole ligands or anionic tridentate tris(pyrazolyl)borate chelates do effect a greater lowering of n(CO) [193]. From past and especially recent insights into Cu NIR structure and spectroscopy, the NIR mechanism consists of: (i) binding of NO 2 to the copper(II) resting enzyme form, most likely via an O- or O;O 0 -coordinated nitrite, (ii) electron transfer from the T1 copper to give a cuprous nitrite, (iii) protonation and loss of water giving a copper-nitrosyl species (formally Cu(I)-(NO1)) which may be side-on bound as found in recent X-ray structures (Figure 31), and release of NO while leaving copper(II) as it all started [194,195].

5.3.2. Nitrous Oxide Reductase In bacterial denitrification and following NIR (nitrite to NO conversion) and then nitric oxide reductase transformation of two moles of NO to give nitrous oxide (N2O), nitrous oxide reductase (N2OR) carries out the terminal step of denitrification: N2O+2 e+2 H1 - N2+H2O. While the reaction is thermodynamically favored, there is a large activation energy Met. Ions Life Sci. 2009, 6, 295–361

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Structure of the CuZ active site of N2OR. Modified from [197].

involved and thus a catalyst is needed. Recent biophysical and especially X-ray crystallographic insights show that each N2OR enzyme monomer unit possesses a ‘purple’ dinuclear electron transfer site called CuA, that consists of a mixed valent center with Cu2(m-Cys)2 core, as also found in cytochrome c oxidase [196]. From here, reducing equivalents are transferred across a subunit interface to the catalytic CuZ center, which consists of a metal cluster containing four copper ions ligated by seven histidine imidazoles, and bridged by an inorganic sulfide ion (Figure 32). It is here that N2O is ‘activated’, reduced and protonated. The CuZ moiety is an extremely novel chemical entity, the first in nature seen to possess an inorganic sulfide together with copper ion; since then a bridging sulfide has been observed in MoCu CODH (Section 3.3). As seen with many copper proteins, the study of small molecule interactions with the accessible CuZ center shed light on the nature of the chemical entity at the catalytic center and/or its function. Here, as relevant to Cu interactions, we note that early studies showed that reaction of certain forms of the enzyme with carbon monoxide leads to activation of the protein, i.e., enhanced reactivity is observed. Riester, Zumft, and Kroneck suggested this may be due to CO acting as a reductant (while itself being oxidized to CO2), which leads to more copper ions in the cluster being in the copper(I) state [198]. We speculate that CO binds to Cu(I) already present or formed in such chemistry, again since CuI-CO interactions are very favorable. In fact, it was more recently demonstrated that the fully reduced all copper(I) CuZ center is indeed the active state in N2O reduction [199,200]. Cyanide also interacts with the CuZ site, perhaps entering as neutral HCN. It inactivates the protein and Haltia and coworkers showed that this treatment leads to the loss of several copper ions from CuZ [196], probably as [CuI(CN)3]2, as mentioned above in Section 3.3. There is a great deal of interest in the mechanism of action of N2OR and how a Cu4S cluster activates nitrous oxide for reduction. Both experiment and theory have recently led to detailed proposals [200]. There is as yet no Met. Ions Life Sci. 2009, 6, 295–361

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example in copper coordination chemistry where copper ion(s) can reduce N2O, although there are ongoing efforts to synthesize copper-sulfide clusters to mimic either the CuZ structure or its reactivity [201–204].

6. COPPER(I) RECOGNITION SITES OR RECEPTORS 6.1. Bacterial Copper Chaperone CusF In Escherichia coli copper homeostasis is in part maintained by the cusCFBA operon, which consists of four proteins (CusC, CusF, CusB, CusA) possessing methionine-rich metal binding pockets [205]. CusCBA forms a multiprotein complex that spans the periplasmic membrane and is responsible for copper efflux processes; CusF is thought to serve as the copper chaperone. Along with spectroscopic inquiries, two recent crystal structures of metallated CusF reveal a unique copper(I)-tryptophan interaction that appears to protect the metal ion from unwanted removal or oxidation [205,206]. The metal binding site of CusF consists of one histidine residue and two methionine residues. Also, upon binding copper(I), the thioether side chains rotate to push the metal ion into a coordination pocket that positions the metal ion 0.5 A˚ out of the trigonal His36-Met47-Met49 plane toward a tryptophan (Trp44) residue. A slight tilt of the Trp44 ring places the Ce3 and Cz3 atoms of the indole ring at 2.67 A˚ and 2.86 A˚ from the copper(I) ion. These Cu-Ctrp distances are longer than reported Z2-CuI-arene distances (see the caption of Figure 33) and along with spectroscopic trends

Figure 33. The CusF binding of copper(I) in a Met2HisTrp coordination environment (middle diagram), and two synthetically derived copper(I) complexes with parene (naphthalene, left; indole ring, right) ligation. The right structure shows copper(I) binding to the indole ring at C2-C3, differing from what is found in CusF. The synthetic complex Cu(I)-Carene bond distances range from 2.13 to 2.41 A˚. Adapted and modified from [207–209]. Met. Ions Life Sci. 2009, 6, 295–361

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found to be similar to non-transition metal systems, they have been designated as cation-p interactions. Cation-p interactions are typically not considered for complexes of transition metals (but see below) because electron sharing between metal d-orbitals and p-orbitals creates covalency [207]. However, supporting the novel copper(I)-tryptophan interaction, titration of copper(I) into solutions of the CusF protein in a stoichiometry of 1:1 resulted in a red shift of the tryptophan absorption at 280 nm and complete quenching of its normal 350 nm emission [205]. Additionally, UVrR data displayed tryptophan mode frequency shifts in the presence of copper(I), similar to previously reported cation-tryptophan p-interactions, suggesting an unconventional cation-p interaction between Trp44 and the copper(I) ion.

6.2. Copper-Ethylene Receptor There is quite strong evidence for the presence of a copper-based ethylene sensor-receptor necessary for proper growth and development in plant cells [210,211]. The ethylene binding receptor (ETR1) of Arabidopsis is believed to have neighboring cysteine (Cys65) and histidine (His64) residues capable of binding copper ions [212]. It is thought that ethylene binding to the copper cofactor results in a change in the coordination environment that is transmitted through the receptor to downstream signaling elements. In fact, there is an extensive literature on the binding of cuprous ion to olefins, including ethylene [213,214]. Such coordination complexes generally show the coordinated ethylene C-C bond to be nearly identical to that of free C2H4. However, 1H and especially 13C NMR spectroscopy are quite useful in demonstrating olefinic-copper(I) interactions and interrogating the bonding in such species.

6.3. Copper Ion in an Olfactory Receptor Site? The olfactory response or sense of smell requires olfactory receptors (Ors) in order to bind and generate a differential response to odorants [215]. In 1978, Crabtree speculated that copper(I) could be a likely candidate to occur at a receptor site [216]. In addition to being able to bind many malodorous species via direct Cu(I)-X interactions, X represents a group 15 (e.g., P or Sb) or group 16 (e.g., S) element, odorants could include carbon-based compounds such as aromatics, olefins, acetylenes, and isocyanides. An odorant metal binding site is believed to be present on the periplasmic domain of the Ors that activates a G-protein binding site, triggering a nerve response that our brain interprets as smell. The structural details of Ors have yet to be determined, but Suslick and coworkers propose a metal Met. Ions Life Sci. 2009, 6, 295–361

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Figure 34. A metal binding site proposed by Suslick and coworkers, here as copper(II) ion, for a consensus amino acid sequence found in the majority of Ors. Adapted and modified from [217] and [215].

coordination geometry predicted to consist of histidine (H), cysteine (C), and glutamate (E) residues (Figure 34) [217]. Approximately 75% of olfactory receptors have a common sequence (HXXC[DE]) on loop regions that extend from the cell membrane; hydrophobic residues are designated as X; D represents aspartate. A synthetic pentapeptide (HAKCE) consisting of histidine (H), alanine (A), lysine (K), cysteine (C), and glutamate (E) was constructed as a putative olfactory binding site and found to have a high affinity for binding copper(II) and other ions, as observed by far-UV CD spectroscopy; metal binding led to a dramatic change typically associated with formation of an a-helical structure. Based on these observations, a ‘‘shuttlecock’’ mechanism was suggested [217]. When an odorant binds to the copper odorant receptor, one of the coordinated amino acids or water molecules is replaced, resulting in protein structural rearrangement. The local charge and steric balance is disrupted and the trans-membrane loop ejects from the membrane triggering a G-protein response (smell). Exposure of the metal bound odorant to extracellular water results in an equilibrium shift, releasing the odorant and allowing the Ors to return to its original state.

7. MISCELLANEOUS 7.1. Bleomycin Bleomycins (BLMs) are a family of glycopeptide-derived antibiotics that require a reduced transition metal (FeII or CuI), dioxygen, and a Met. Ions Life Sci. 2009, 6, 295–361

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Figure 35. Schematic of the proposed mechanisms (right) for generation of active DNA damaging bleomycins; modified from [218,222]. The X-ray structure to the left is a conformational comparison of copper(II)-BLM and cobalt(III)-BLM [218,222].

one-electron reductant to effect single- or double-stranded cleavage of the sugar-phosphate backbone of DNA [218]. BLMs are isolated from Streptomyces verticillus in the copper(II)-bound form and have developed into an essential anticancer agent [219]. Current chemotherapy protocols involving this natural-product-derivative have been used to treat Hodgkin’s lymphoma, testicular and ovarian cancer, as well as carcinomas of the head, skin, and neck, Figure 35 [218,220,221]. The proposed mechanism for the generation of activated bleomycin involves metal-free BLM quickly picking up copper(II) ions from blood plasma to be transported into the cell. CuII-BLM must then be reduced by one electron to CuI-BLM at which point intracellular exchange for iron(II) occurs. Reaction of the newly formed FeII-BLM with dioxygen then results in the activated bleomycin derivative, BLM-FeIII-OOH [223]. As a result, the copper(II) BLM adduct has been labeled by some researchers as a prodrug that simply delivers BLM into the cell nucleus at which point copper(I) BLM becomes a spectator to the DNA damaging ability of the iron(III) hydroperoxo species [221]. However, a more biologically significant role for CuI-BLM has also been suggested. Met. Ions Life Sci. 2009, 6, 295–361

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Hecht, Oppenheimer, and coworkers investigated the intracellular exchange process of copper(I) for iron(II) through two-dimensional 1H-13C correlated spectroscopy [224]. CuI-BLM was determined to be in rapid equilibrium with copper(I) ions and metal-free bleomycin. The exchange process decreased in the presence of carbon monoxide due to formation of BLM-CuI-CO. Therefore, chemical shift assignments could be confirmed for metal-free BLM, CuI-BLM, and BLM-CuI-CO. The more significant discovery was that DNA degradation by CuI-BLM occurred independently of iron(II). In a later study it was also discovered that DNA damage by bleomycin in the presence of both iron(II) and copper(I) was greater than with either metal alone [225]. Also, the NMR data suggested that CuI-BLM is structurally distinct from FeII-BLM. Since this report, Lehmann has suggested that the geometry upon metal chelation is important for BLM activity [221]. The first X-ray crystal structure of a copper(II)-complexed form of bleomycin, designated CuII-BmA2, bound to a bleomycin resistance determinant (BLMA) from S. verticillus, was reported by Matoba and coworkers [222]. BmA2 refers to a form of BLM in which the bithiazole tail (DNA recognition domain) is coordinated to a g-aminopropyl dimethylsulfonium moiety, (see Figure 36). CuII-BmA2 exhibits ligation from five N-donors in a pyramidal geometry with the primary amine of the b-aminoalanine moiety

Figure 36. Structure of BLM with the five N donors responsible for metal chelation in bold. The four functional domains of the structure are separated and labeled (see text for more detail). This BLM structure diagram has been modified from [218,226]. Met. Ions Life Sci. 2009, 6, 295–361

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in the axial position. The bithiazole tail is in an extended conformation in the copper(II) X-ray structure as well as in the metal-free form of BmA2 bound to BLMA. Based on two-dimensional NMR experiments, Lehmann has recently suggested that a similar geometry in the copper(I)-BLM adduct prevents DNA degradation and instead favors intracellular transformation of the copper(I) species to the iron(II) adduct [221]. The actual form of BLM inside the cell has yet to be established. Various metal ions such as iron, copper, cobalt, manganese, nickel, ruthenium, vanadium, and zinc are known to bind to BLM and lead to DNA degradation [226]. Actually, studies have shown that the level of cytotoxicity is not affected by the form of the metallobleomycin, whether zinc, copper, nickel, or iron [218]. As a result, the in vivo form of BLM is still under debate.

7.2. Copper-Alkyl Complexes from Biologically Derived Carbon Radicals Meyerstein and coworkers have carried out extensive studies on reactions of aqueous radicals with transition metal ions, including copper [227,228]. Depending on the other copper-ligand(s) present, either Cu(I) or Cu(II) can lead to transient Cu-alkyl complexes via the reaction, Cun+Rd - Cun11(R). Such reactions are very fast and would very well compete with any other reactions of alkyl radicals (for example with other biomolecules). The major route for decomposition when the initial reactant is cuprous ion would be CuC heterolysis: CuII(R)+H2O - CuI+RH+OH. When starting with cupric ion, typical reactions observed are: CuIII-CH3 - CuI(aq)+CH3OH+H1, or for example, (glycinate)2CuIII-CH3 - (glycinate)2CuIII+CH4+OH. Reactions leading to CuIII could be biologically harmful, since this is a very strong oxidant. However, Meyerstein points out that the hypothesis that copper (or other) ion reactions with alkyl radicals leading to deleterious processes needs to be confirmed in actual biological systems.

8. GENERAL CONCLUSIONS In fact, copper-carbon bonds are formed in biological systems, especially in the extensive use of carbon monoxide and cyanide as spectroscopic or chemical probe ligands for either cuprous or cupric ion protein active sites. For cuprous species, observation of copper(I)-carbonyl CO stretching frequencies gives valuable comparative information about the copper local environment. For example, carbonmonoxy protein derivatives possess revealing spectroscopic emission properties that provide insight about the basic coordination properties of these novel chemical entities. In some cases, Met. Ions Life Sci. 2009, 6, 295–361

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isocyanides have also been utilized for similar purposes. Cyanide can extract copper ion from protein active sites, or its binding can allow for spectroscopic (e.g., UV-vis, CD, EPR) interrogation to provide information about coordination geometries in the solution state. More recently, it has come to light that in several cases, carbon-containing moieties are intimately involved with copper protein function: Carbon monoxide is the actual enzyme substrate in the molybdenumcopper containing MoCuCODH where it binds to a copper(I) protein site and is then oxidatively transformed to carbon dioxide. Plants apparently employ a copper(I) receptor to detect (through binding) the presence of ethylene, which is a growth hormone. In CusF, a p-arene fragment of a protein tryptophan residue contributes to selectivity and coordination of copper(I) ions in this copper(I) chaperone protein. It is speculated that copper ion could be the basis of some odorant receptor sites, based on its ability to bind carbonaceous odorants, as well as many other heteroatom possessing molecules. If one reflects broadly, it would seem that the sub-field of biologically relevant copper-carbon interactions is growing. We speculate that it may be found in the future that biological receptors which bind CO as a messenger may be copper(I)-based. A good deal of literature indicates CO is a neural messenger, involved in cell signaling, possibly also (like NO) a physiological regulator of cyclic guanosine monophosphate [229,230]. While heme-based sensors/receptors for CO have been discussed, copper(I) sites may also function similarly. In certain cases it is known that CO possesses beneficial biological effects as a protein inhibitor [231,232]. Perhaps copper-CO compounds may be developed as drug sources of carbon monoxide; with respect to other transition metal-carbonyls, this is an area being currently investigated [233,234]. We note that another use of copper-carbon interactions has been with synthetically derived 11C-labeled CN–-copper(I) for the purpose of radiolabeling biologically active ligands or for use in positron emission tomography [235,236]. Finally, a related and perhaps somewhat obvious use of copper(I) compounds would be to act as biosensors/monitors of the toxic cyanide or carbon monoxide molecules.

ACKNOWLEDGMENTS The authors are very grateful to the National Institutes of Health (USA) for support in writing of this article, and for carrying out the copper or heme/ copper chemistry mentioned here. Met. Ions Life Sci. 2009, 6, 295–361

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ABBREVIATIONS AGAO AO ATP Az BLM BLMA BmA2 CAO CbO CcO CD CODH Cp CuZn SOD DFT DIMPI DbM EPR ET ETR1 EXAFS FTIR GO Hc HcCO HCO IbCO IPI IR Lc Mb MCO 1-MeIm 1,2-Me2Im MePY2 MLCT MoCu CODH mtp MWC NHE NIR

Arthrobacter globiformis amine oxidase ascorbate oxidase adenosine 5 0 -triphosphate azurin bleomycin bleomycin resistance determinant bleomycin A2 copper amine oxidase bo3 oxidase from E. coli cytochrome c oxidase circular dichroism carbon monoxide dehydogenase ceruloplasmin copper-zinc superoxide dismutase density functional theory 2,6-dimethylphenyl isocyanide dopamine b-monooxygenase electron paramagnetic resonance electron transfer ethylene binding receptor extended X-ray absorption fine structure Fourier transform infrared galactose oxidase hemocyanin carbon monoxy-hemocyanin heme-copper oxidase Ipomoea catechol oxidase isopropyl isocyanide infrared laccase myoglobin multi-copper oxidase 1-methylimidazole 1,2-dimethylimidazole bis(2-pyridylethyl)methylamine metal-to-ligand charge transfer molybdenum-copper carbon monoxide dehyrogenase metallopterin Monod-Wyman-Changeux normal hydrogen electrode nitrite reductase Met. Ions Life Sci. 2009, 6, 295–361

352

NMe2-tmpa NMR NMRD NOR Ors PAC PBD Pc PHM PHMcc PY1 RbCcO RNC R-PYAN rR SOD Solv t-BuNC THF tmpa TPQ UVrR XAS

LUCAS and KARLIN

tris(4-dimethylaminopyrid-2-ylmethyl)amine nuclear magnetic resonance spectroscopy nuclear magnetic resonance dispersion nitric oxide reductase olfactory receptors photoacoustic calorimetry photothermal beam deflection plastocyanin peptidylglycine a-hydroxylating monooxygenase catalytic core of peptidylglycine a-hydroxylating monooxygenase bis(2-pyridylmethyl)amine cytochrome c oxidase from Rhodobacter sphaeroides isocyanide N-[2-(4-R-pyridin-2-yl)-ethyl]-N,N 0 ,N 0 -trimethyl-propane-1,3-diamine; R ¼ NMe2, OMe, H, and Cl resonance Raman superoxide dismutase solvent tert-butyl isocyanide tetrahydrofuran tris(pyrid-2-ylmethyl)amine 2,4,5-trihydroxylphenylalanine quinone UV resonance Raman X-ray absorption spectroscopy

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10 Interaction of Cyanide with Enzymes Containing Vanadium, Manganese, Non-Heme Iron, and Zinc Martha E. Sosa-Torres a and Peter M. H. Kroneck b a

Facultad de Quı´ mica, Universidad Nacional Autońoma de Me´xico, Ciudad Universitaria, Coyoacań, 04510, D.F. Me´xico, Me´xico <[email protected]> b Fachbereich Biologie, Universita¨t Konstanz, D-78457 Konstanz, Germany

ABSTRACT 1. INTRODUCTION 1.1. Cyanide. Prebiotic Substrate and Inhibitor 1.2. Cyanide. Ligand and Probe for Transition Metal Sites 2. VANADIUM ENZYMES 2.1. Vanadium in Biology. Structures and Functions 2.1.1. Vanadium Haloperoxidases 2.1.2. Vanadium Nitrogenase 2.2. Vanadium Enzymes and Their Interaction with Cyanide 2.2.1. Vanadium Haloperoxidases 2.2.2. Vanadium Nitrogenase 3. MANGANESE ENZYMES 3.1. Manganese in Biology. Structures and Functions 3.1.1. Manganese Superoxide Dismutase 3.1.2. Manganese Catalases 3.2. Manganese Enzymes and Their Interaction with Cyanide 3.2.1. Manganese Superoxide Dismutase 3.2.2. Manganese Catalase Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00363

364 364 365 366 368 368 368 370 372 372 372 373 373 374 375 376 376 377

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SOSA-TORRES and KRONECK

4. NON-HEME IRON ENZYMES 4.1. Non-Heme Iron Enzymes. Structures and Functions 4.1.1. Enzymes Carrying the 2-His-1-Carboxylate Facial Triad Structural Motif and Variants 4.1.2. Protocatechuate 3,4-Dioxygenase 4.1.3. The Non-Heme Iron Center of the Oxygen-Evolving Complex of Photosystem II 4.2. Non-Heme Iron Enzymes and Their Interaction with Cyanide 4.2.1. Enzymes Carrying the 2-His-1-Carboxylate Facial Triad Structural Motif and Variants 4.2.2. Protocatechuate 3,4-Dioxygenase 4.2.3. The Non-Heme Iron Center of the Oxygen-Evolving Complex of Photosystem II 5. ZINC ENZYMES 5.1. Zinc in Biology. Structures and Functions 5.1.1. Zinc Carbonic Anhydrase 5.1.2. Zinc Hydrolases 5.2. Zinc Enzymes and Their Interaction with Cyanide 5.2.1. Zinc Carbonic Anhydrase 5.2.2. Zinc Hydrolases 6. CONCLUSIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

377 377 378 380 380 381 381 382 382 383 383 384 386 387 387 388 388 388 388 389

ABSTRACT: Since the early discovery of Prussian Blue, cyano transition metal complexes have played a fundamental role in coordination chemistry. They represent important compounds with fascinating chemical and physical properties which turn them into valuable tools for both chemists and biologists. HCN as a precursor in prebiotic chemistry has gained interest in view of its polymers being involved in the formation of amino acids, purines, and orotic acid, a biosynthetic precursor of uracil. Clearly, the rapid formation of adenine by aqueous polymerization of HCN is one of the key discoveries in these experiments. The cyanide anion is usually toxic for most aerobic organisms because of its inhibitory effects on respiratory enzymes, but as a substrate it is an important source of carbon and nitrogen for microorganisms, fungi and plants. Most interestingly, the cyanide anion is a ligand of important metal-dependent biomolecules, such as the hydrogenases and the cobalt site in vitamin B12. KEYWORDS: cyanide inhibition cyano complexes manganese superoxide dismutase non-heme iron center Prussian Blue vanadium nitrogenase

1. INTRODUCTION The cyanide anion (CN) is well known to be highly toxic to living cells and usually causes inhibition of growth, because of (i) tight coordination to Met. Ions Life Sci. 2009, 6, 363–393

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metal centers of respiratory metallo-enzymes, such as cytochrome c oxidase, (ii) reaction with keto compounds to form cyanohydrin derivatives of enzyme substrates, and (iii) reaction with Schiff-base intermediates to form stable nitrile derivatives. Despite its toxicity, many microorganisms have developed cyanide-resistant respiratory systems to enable them to survive and grow in the presence of cyanide [1]. Some microorganisms are able to use cyanide as a source of nitrogen, or even carbon, for growth. Early on, the biochemistry of cyanide attracted the interest of numerous researchers, and we recommend two books for introduction: (i) Cyanide in Biology [2], and (ii) Cyanide Compounds in Biology [3]. As the study of cyano metal complexes may be said to have begun with the accidental discovery of Prussian Blue, Fe4[Fe(CN)6]3, by the Berlin painter Diesbach more than 300 years ago [4], we also suggest the book by Sharpe entitled The Chemistry of Cyano Complexes of the Transition Metals [5] as much of our understanding of coordination chemistry can be attributed to research on cyano metal complexes. A renaissance of Prussian Blue chemistry and its transition metal analogues has occurred in the past decade because of exciting properties of cyanide materials [6]. Furthermore, the Handbook on Metalloproteins, [7], and the Handbook of Metalloproteins [8,9] summarize valuable information on structural and biochemical properties of metalloproteins. Perhaps the most prominent example of a biological cyano complex is vitamin B12 which is discussed at length by Kraütler in Chapter 1. Note that several microorganisms are able to synthesize the complex B12 structure including variations of the nucleotide function. In general, cobalt-corrins have been proposed to be structural and functional remnants of early forms of life. Most likely, central metabolic processes depended on organometallic compounds employing either cobalt or nickel centers [10]. Recently, a rather surprising feature came from the characterization of hydrogenases applying Fourier transform infrared spectroscopy. These spectroscopic investigations revealed that the metal atoms at the active site cluster (Ni and Fe) were surrounded by carbon monoxide and cyanide [11]. Although such features are not unusual in laboratory-produced catalysts, they were hard to fathom in an enzyme [12]. Clearly, the synthesis of the CN and CO ligands of the hydrogenases poses intriguing questions because of their toxicity in the free state, and in particular, because of addition of CO and CN to the metal reflects organometallic chemistry unprecedented in biology [13]. Enzymethiocarbamate and thiocyanate complexes are formed on the biosynthetic pathway, the latter acting as the cyanide donor to the iron center.

1.1. Cyanide. Prebiotic Substrate and Inhibitor Cyanide might have played a fundamental role in the evolution of life on earth [14]. In the context of prebiotic chemistry, the acid/base hydrolysis of Met. Ions Life Sci. 2009, 6, 363–393

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HCN and derivatives including cyanoacetylene and the HCN tetramer has been explored extensively [15–17]. It represents a valuable form of nitrogen for microorganisms, fungi and plants. Although several organisms synthesize cyanide, a greater number are capable of cyanide biodegradation [18,19]. The existence of these pathways has led to the development of biotechnological processes to degrade cyanide compounds in industrial waste streams, originating from activities such as petrochemical refining, production of organic chemicals and polymers, and metal mining and processing. Representative examples of chemical reactions for the transformation of cyanide and cyanide compounds are listed below (reactions 1–5). Cyanide hydratase HCN þ H2 O ! HCONH2

ð1Þ

HCN þ 2H2 O ! HCOOH þ NH3

ð2Þ

RCN þ 2H2 O ! RCOOH þ NH3

ð3Þ

Cyanidase Nitrilase

Cyanide monooxygenase HCN þ O2 þ Hþ þ NADðPÞH ! HOCN þ NADðPÞþ þ H2 O Rhodanese

2 CN þ S2 O2 3 ! SCN þ SO3

ð4Þ ð5Þ

On the other hand, cyanide, like azide ðN 3 Þ and carbon monoxide (CO), has been used as a valuable tool to study electron transfer processes in respiratory chains of numerous organisms. Hereby, cyanide acts as an irreversible inhibitor and binds to metal sites, with cytochrome c oxidase being perhaps the most prominent example [20]. Using infrared and visible Soret spectra the binding of cyanide to both iron and copper sites of the enzyme in its different redox states could be demonstrated.

1.2. Cyanide. Ligand and Probe for Transition Metal Sites Cyanide is one of the most common and longest-known ligands in coordination chemistry. Cyanide complexes are often compared to metal carbonyls, and both CN and CO can function as p-acid ligands. However, the cyanide ion possesses a negative charge, and can form very strong s-bonds with metal ions. With very few exceptions, all cyanide complexes display metal-carbon interactions, a characteristic of organometallic compounds [21,22]. The s-donor capacity of CN allows it to stabilize transition metals in high oxidation states. In addition, it can function as a p-acid ligand, by accepting electron density from a filled metal d orbital into an empty Met. Ions Life Sci. 2009, 6, 363–393


367

antibonding orbital of the carbon-nitrogen bond. This allows CN to stabilize transition metals with low oxidation numbers. The binding properties of CN make it a very strong-field ligand, actually one of the highest in the spectrochemical series. The cyanide anion also exhibits a strong trans effect and a large nephelauxetic effect, indicative of the high degree of covalency in the metal-cyanide bond [23] (Figure 1). Consequently, the cyanide anion proved to be an excellent tool to probe Fe(III) heme centers by spectroscopic techniques, especially 13C NMR and EPR. Cyanide shows little affinity for Fe(II) myoglobin and hemoglobin, but binds favorably to Fe(III) hemoproteins. It is frequently chosen as a strong-field ligand to ensure conversion of Fe(III) porphyrins to the lowspin state. Thus, in analogy to carbon monoxide ligation by reduced hemoproteins, CN can provide an axial ligand probe for the protein in the Fe(III) state. These investigations allow to develop a well-resolved picture of the electronic properties of the metal site [24,25].

Antibonding mostly metal E

4p 4s π* eg 3d

Antibonding mostly ligand Antibonding mostly metal

t2g Bonding mostly metal

π

Bonding mostly ligand

Mn+

M(CN)6n-6

CN-

Figure 1. Schematic molecular orbital energy level diagram for a complex [M(CN)6]n6. Met. Ions Life Sci. 2009, 6, 363–393

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Cyanide was also developed into a useful ligand-directed probe of the coordination site of Cu,Zn-superoxide dismutase employing both infrared and Raman spectroscopy [26]. Finally, cyanide has been successfully employed to inactivate molybdoenzymes, such as xanthine oxidase or aldehyde oxidase, producing the desulfo form of the active site. The so-called cyanolyzable sulfur bound to the molybdenum center reacts with cyanide to form thiocyanate, SCN, which can be quantitatively determined [27].

2. VANADIUM ENZYMES 2.1. Vanadium in Biology. Structures and Functions The biological function of vanadium is well established and the chemistry of the element is gaining considerable interest. The metal is an inherent part of enzymatic active sites, prominent examples are the vanadium-containing haloperoxidases and vanadium nitrogenase [28]. For both types of Vdependent enzymes there exist functional analogues in nature which are either more widely spread or more efficient, for example, the heme-containing haloperoxidases and the conventional Mo-dependent nitrogenases, respectively. One might ask how these enzyme systems evolved, and in particular, whether the vanadium-containing enzymes known today are retained functional analogues, which withstood evolutionary forces. The widespread physiological effects of vanadium are mainly attributed to the similarity of the vanadate(V) ions and phosphate ions ðPO3 4 Þ. Note that, depending on pH, also important differences exist between theses two anions with regard to charge, redox and coordination properties [28,29].

2.1.1. Vanadium Haloperoxidases Vanadium-containing haloperoxidases are enzymes isolated primarily from marine algae, although they also have been purified from other organisms [30]. These enzymes catalyze the two-electron oxidation of halide ions (X) to the corresponding hypohalous acids, HOX (equation 6). HOX can further react with a broad range of acceptors to form a diversity of halogenated compounds [31]. Generally, the haloperoxidases are named after the most X þ Hþ þ H2 O2 ! O2 þ H2 O þ HOX

ð6Þ

electronegative halide they can oxidize, thus vanadium chloroperoxidase, in addition to chloride, will also transform bromide. Recently, two different Met. Ions Life Sci. 2009, 6, 363–393


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haloperoxidases, one specific for the oxidation of iodide, and the second that can oxidize both iodide and bromide, were obtained from the brown alga Laminaria digitata. The iodoperoxidase activity was significantly more efficient than the bromoperoxidase fraction in the oxidation of iodide. The two enzymes differed markedly in their molecular masses, trypsin digestion profiles, and immunological characteristics [32]. One of the best characterized V-dependent chloroperoxidases is the fungal enzyme isolated and purified from Curvularia inaequalis [33]. Its X-ray structure (2.1 A˚), in complex with azide, reveals that the ligand coordinates directly to the vanadium center. Furthermore, three non-protein oxygen ligands and one histidine nitrogen are bound. In the native state, vanadium is bound as hydrogen vanadate(V) in a trigonal bipyramidal geometry, with the metal coordinated to three oxygens in the equatorial plane, and HO (from water) and a nitrogen of histidine at the apical positions. The vanadium-containing bromoperoxidase from the seaweed Ascophyllum nodosum shows high similarities in the regions of the metal binding site, with all hydrogen vanadate(V) interacting residues conserved except for Lys353 which has been replaced by an asparagine. An interesting observation results from a trapped phosphate intermediate in the crystal structure (1.5 A˚) of vanadium-free apochloroperoxidase from Curvularia inaequalis which catalyzes a dephosphorylation reaction [34]. Since the chloroperoxidase is functionally and evolutionary related to several acid phosphatases including human glucose-6-phosphatase and a group of membrane-bound lipid phosphatases, the structure may help to understand the mechanism of action of these enzymes as well. The trapped intermediate is bound to the active site as metaphosphate, PO 3 , with its phosphorus atom covalently attached to the nitrogen atom of His496. An apical water molecule is within hydrogen-bonding distance to the phosphorus atom, and it is in a perfect position for a nucleophilic attack on the metaphosphate-histidine intermediate to form the inorganic phosphate. Recently, the 3D structure of a vanadium-dependent bromoperoxidase dodecamer from the red algae Corallina officinalis has been determined at 2.3 A˚ resolution, with each subunit made up of 595 amino acid residues [35]. A cavity is formed by the N terminus of each subunit in the center of the dodecamer. The subunit fold and dimer organization of the bromoperoxidase are similar to those of the dimeric enzyme from the brown algae Ascophyllum nodosum, with which it shares 33% sequence identity. The different oligomeric states of the two algal enzymes seems to reflect separate mechanisms of adaptation to harsh environmental conditions and/or to chemically active substrates and products. The residues involved in the vanadate binding are conserved between the two bromoperoxidases and the vanadium chloroperoxidase from Curvularia inaequalis. However, most of the other residues forming the active-site cavity are different in the three Met. Ions Life Sci. 2009, 6, 363–393

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enzymes, which reflects differences in the substrate specificity and stereoselectivity of the reaction. The vanadate is directly linked to the protein only through the axially bound histidine residue, and is embedded in the protein through an extensive hydrogen-bonding network.

2.1.2. Vanadium Nitrogenase Nitrogenase, the microbial enzyme catalyzing biological nitrogen fixation accounts for the cycling of at least 9 1010 kg of nitrogen fixed annually [36]. All N2-fixing organisms (diazotrophs) which have been investigated have a nitrogenase system based on Mo and Fe. In addition, it is now known that some organisms have alternative nitrogenases based on V and Fe, or apparently Fe alone [37]. Under optimal conditions Mo-nitrogenase catalyzes the ATP-dependent reduction of dinitrogen to ammonia and protons to dihydrogen employing a two-component enzyme machinery (reaction 7). N2 þ 8Hþ þ 8e þ 16MgATP ! 2NH3 þ H2 þ 16MgADP þ 16Pi

ð7Þ

Vanadium was shown unequivocally to have a role in biological nitrogen fixation, when V-containing nitrogenases were isolated from mutant strains unable to synthesize Mo-nitrogenase [38,39]. The reality of a Mo-independent route for nitrogen fixation was established beyond question when the structural genes encoding Mo-nitrogenase in Azotobacter vinelandii were specifically removed and the resulting deletion mutant strains shown to be able to fix N2 in Mo-deficient medium [40]. Subsequently, V-nitrogenases were purified from strains of Azotobacter chroococcum and Azotobacter vinelandii [37]. The VFe protein contains P cluster redox centers and a catalytic FeV cofactor, in which vanadium is in a polynuclear cluster with iron, sulfur, and homocitrate with a chemical environment similar to molybdenum in MoFe proteins [41]. The crystal structures of both individual proteins of Mo-nitrogenase and the putative ADP-AlF4 transition state complex of the two proteins have been determined [42,43]. The Fe protein is a dimer which has a single [4Fe-4S] center ligated at the subunit interface and two nucleotide-binding sites, one on each subunit. The structures of the MoFe proteins have revealed an a2b2-subunit structure in which each dimeric ab-subunit pair binds the [8Fe-7S] P-cluster positioned at the subunit interface and the [7Fe-9S-homocitrate] FeMo cofactor center within the a-subunit. This information, together with comparative spectroscopic data enabled a more meaningful interpretation of data obtained for V-nitrogenase, for which the structure has yet to come (Figure 2) [44]. Met. Ions Life Sci. 2009, 6, 363–393


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Figure 2. Stick-and-ball model of the nitrogenase FeMo-cofactor including the m6interstitial atom (blue) in the center of the Fe-S moiety (Fe, S, and Mo are shown as grey, yellow and orange spheres) [45]. In the current model of the FeV-cofactor, vanadium would replace molybdenum [46].

The redox centers of V-nitrogenase have been investigated by EPR, MCD, Mo¨ssbauer, and X-ray absorption spectroscopies [47–50]. The spectroscopic data for VFe proteins are fully consistent with the presence of redox centers very similar to those of Mo-nitrogenase. An indication of the similarity of these systems is the ability of the components of V-nitrogenase to form fully functional hybrid nitrogenases with components of the Mo-nitrogenase [47]. V-nitrogenase of Azotobacter chroococcum has been reported to consume more ATP and to produce more dihydrogen by comparison with Monitrogenase, the reasons and possible significance of these differences remain currently unclear [28]. The V-nitrogenase from Azotobacter chroococcum produces ammonia and traces of hydrazine [51], and V-nitrogenase also Met. Ions Life Sci. 2009, 6, 363–393

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exhibits an alkyne reductase activity as well as an isocyanide reductase activity [52–54].

2.2. Vanadium Enzymes and Their Interaction with Cyanide 2.2.1. Vanadium Haloperoxidases Vanadium haloperoxidases catalyze the oxidation of halide anions, Cl, Br and I, by hydrogen peroxide (reaction 6). Recently, Pecoraro and coworkers suggested that the V-haloperoxidases may actually be peroxidases and that the well-known halogenation reactions may simply reflect the abundance of chloride in the marine environment [55]. The potentials for the oxidation of I, Br and Cl span a range of 0.53–1.36 V (versus NHE), the potentials for the oxidation of the pseudohalide anions, such as CN, OCN, or SCN, fall within or below this range, suggesting the vanadium haloperoxidases may also catalyze the transformation of these compounds. Indeed, cyanide (Eo ¼ +0.375 V) acts as an inhibitor of V-bromoperoxidase isolated from the marine alga Ascophyllum nodosum through preferential oxidation of CN. Similarly, thiocyanate (Eo ¼ +0.77 V) was reported to inhibit bromide peroxidation through preferential oxidation of thiocyanate over bromide. 13C NMR studies of the oxidation of K13CN by H2O2 catalyzed by V-bromoperoxidase showed the formation of several oxidized thiocyanate species, including the putative (unstable) dithiocyanate ether, hypothiocyanate, thiooxime, and bicarbonate [46,56]. Bromine K-edge EXAFS experiments on samples containing bromide and V-peroxidase, in buffer pH 8, were carried out, with biomimetic vanadium compounds carrying Br-V, Br-C(aliphatic), and Br-C(aromatic) bonds, as reference. From these studies it appears that bromide does not coordinate to the vanadium center of the peroxidase, but binds covalently to carbon, with an active site serine as possible candidate. In this series of experiments, two oxovanadium complexes, [VO(H2O)2(sal-L-Leu)] and [VO(H2O)2(5-Br-sal-Gly)] have been structurally characterized [46]. These complexes contain the water ligands in cis and trans positions to the oxo group, at V-OH2 distances ranging from 2.008 to 2.228 A˚, and were used to model the apical electron density feature observed in the structures of fungal and algal V-haloperoxidases.

2.2.2. Vanadium Nitrogenase Mo-nitrogenase is able to reduce both CN and N 3 to NH3. Because V-nitrogenase has been shown to release hydrazine, N2H4, in the course of Met. Ions Life Sci. 2009, 6, 363–393


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N2 reduction it was chosen to investigate the conversion of cyanide and azide. Sensitive assay procedures were developed to monitor the production of either HCHO or CH3OH from HCN. Like Mo-nitrogenase, the vanadium enzyme suffered electron flux inhibition when interacting with CN, but in contrast to Mo-nitrogenase, MgATP hydrolysis was also inhibited by CN in the case of V-nitrogenase. At high concentrations of cyanide, the vanadium enzyme directed a significant percentage of electrons into the production of excess NH3. Under these experimental conditions, a substantial amount of formaldehyde (HCHO) but not methanol (CH3OH) was detected for the first time [57]. Azide inhibited both the total electron flux as well as MgATP hydrolysis in the case of V-nitrogenase but not Monitrogenase. V-nitrogenase, unlike Mo-nitrogenase, revealed no preference between the two electron reduction to N2-plus-NH3 and the six-electron reduction to N2H4-plus-NH3. V-nitrogenase formed more excess NH3, but reduction of the N2 produced by the two electron reduction of N 3 . Unlike Mo-nitrogenase, CO could not completely eliminate either cyanide or azide reduction by V-nitrogenase. CO did, however, eliminate the inhibition of both electron flux and MgATP hydrolysis by CN, but not that caused by azide. These different responses to CO suggest different sites or modes of interaction for these two substrates with V-nitrogenase.

3. MANGANESE ENZYMES 3.1. Manganese in Biology. Structures and Functions Manganese can play many roles in biology ranging from acting as a simple Lewis acid catalyst to being an element that can transverse several oxidation states to carry out water oxidation [58,59]. Manganese is an essential constituent of the tetranuclear Mn cluster that is involved in dioxygen production in photosynthetic plants, algae, and cyanobacteria. Water splitting is driven by the membrane pigment-complex known as photosystem II-water oxidizing complex (PSII-WOC), or oxygen-evolving complex (PSII-OEC). The electrons and protons are ultimately used to store energy in the form of ATP and to reduce CO2 to carbohydrate. The complex is directly involved in the oxidation of water, it carries four Mn ions embedded in the protein matrix [58]. Crystallographic investigations of cyanobacterial PSII-OEC provided several medium-resolution structures (3.8–3.2 A˚) that explained important features of the protein matrix and cofactors, but did not produce a highly resolved picture of the complex [60–63]. The most complete cyanobacterial photosystem II structure reported so far (3 A˚) showed locations of and interactions between 20 protein subunits and 77 cofactors per monomer. Assignment of 11 b-carotenes yielded insights into electron and Met. Ions Life Sci. 2009, 6, 363–393

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energy transfer and photo-protection mechanisms in the reaction center and antenna subunits, and the structure provided further information about the Mn4Ca cluster, where oxidation of water takes place [64]. Nevertheless, many questions remain to be solved about the structure and mode of action of PSII-OEC as discussed most recently [65]. Manganese is the cofactor for superoxide dismutases, catalases and some peroxidases, which are all used for the detoxification of reactive oxygen species (ROS). An important property of manganese in its 2+ oxidation state, which has important biochemical consequences, is that it is a close but not exact surrogate of Mg21. Mn21 with its relatively similar ionic radius can readily exchange with Mg21 in most structural environments, and exhibits much of the labile, octahedral coordination chemistry. However, it can more easily accommodate the distortions in coordination geometry in progressing from the substrate-bound to the transition state and to the bound product. Consequently, Mn21 in the active site of a Mg21-enzyme often results in improved enzyme efficacy [10].

3.1.1. Manganese Superoxide Dismutase There are relatively few characterized proteins having mononuclear Mn sites. By far the best characterized mononuclear site is that in Mn-superoxide dismutase which catalyzes the disproportionation of superoxide ðO 2 Þ to hydrogen peroxide and dioxygen (reaction 8). The overall reaction is a redox þ 2O 2 þ 2H ! H2 O2 þ O2

ð8Þ

process that involves the alternate oxidation and reduction of the catalytic active site metal. In the first step, superoxide binds to the resting Mn(III) enyzme, and is oxidized to dioxygen, with Mn(III) being reduced to Mn(II). In the second step, a second molecule of superoxide binds to the Mn(II)enzyme, and is reduced to hydrogen peroxide, with Mn(II) being reoxidized to the Mn(III) state. There are four different types of superoxide dismutases known, carrying Cu and Zn, Fe, Mn, or Ni in the active site. In prokaryotes, Mn-superoxide dismutase is most commonly found as a dimer in which two monomers come together at a highly conserved interface. The eukaryotic enzyme is usually tetrameric, formed when two of the prokaryotic-like dimers come together to form a dimer of dimers. Each subunit in the oligomer has one metalbinding active site, found at the junction of the two domains of which the monomer is composed. The crystal structures of several Mn-superoxide dismutases including the human enzyme have been determined [66,67]. Met. Ions Life Sci. 2009, 6, 363–393


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The metal site has an approximately trigonal bypyramidal structure with an equatorial plane of two histidine imidazoles and one aspartate carboxylate, and axially coordinated solvent and imidazole.

3.1.2. Manganese Catalases The Mn-catalases possess a dinuclear Mn active site and catalyse the disproportionation of hydrogen peroxide (reaction 9). Catalases play an important protective role by converting toxic hydrogen peroxide into dioxygen and water. In contrast to their heme-containing counterparts, which are ubiquitous in aerobic organisms, a broad range of microorganisms, living in anoxic, or close to anoxic environments, have Mn-catalases. A most notable property of this type of catalase appears to be its insensitivity to the classical heme poisons, cyanide and azide [68]. 2H2 O2 ! 2H2 O þ O2

ð9Þ

Most of the work on Mn-catalases has focused on the enzymes isolated from Lactobacillus plantarum and Thermus thermophilus. The X-ray structure of Mn-catalase from Lactobacillus plantarum [69] revealed the dimanganese active site (Figure 3). The oxidized [Mn(III)Mn(III)] cluster is

Figure 3.

The active site of Lactobacillus plantarum Mn-catalase [69]. Met. Ions Life Sci. 2009, 6, 363–393

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bridged by two solvent molecules (oxo and hydroxo, respectively) together with a m-1,3-bridging glutamate carboxylate and is embedded into a network of hydrogen bonds involving a tyrosine residue (Tyr42). Spectroscopic and structural studies indicate that disruption of the hydrogen-bonded network significantly perturbs the active site in the Y42F variant. This variant has less than 5% of the catalase activity and much higher Km for H2O2 (E1.4 mM) at neutral pH than the wild-type enzyme, although the activity is slightly restored at high pH. The occurrence of m-1,3-bridging carboxylates appears to be a universal feature of bimetalloproteins and complexes which perform two-electron or multi-electron chemistry. Bridging carboxylates probably serve a functional role beyond merely that of a passive structural bridge to bring the metals together. The dinuclear Mn center carries out a two-electron catalytic cycle, interconverting between reduced [Mn(II)Mn(II)] and oxidized [Mn(III)Mn(III)] states during turnover [70].

3.2. Manganese Enzymes and Their Interaction with Cyanide 3.2.1. Manganese Superoxide Dismutase Mn-superoxide dismutases are usually insensitive towards inhibition by cyanide, in fact that is the way of recognizing them. About three deacades ago, in studies on micronutrient interactions in plants, leaf extracts of Pisum sativum showed the presence of three electrophoretically distinct superoxide dismutases, two of which were inhibited by cyanide while the third one was insensitive towards cyanide [71]. The CN-resistant activity, as judged by its dependence on manganese, appeared to be rather a Mn-superoxide dismutase than a Fe-superoxide dismutase. Azide ðN 3 Þ, on the other hand, proved to be a competitive inhibitor of Mn-SOD, and it is believed to bind analogously to superoxide. As expected, the crystal structure of the azide derivative of Mn-SOD from Thermus thermophilus shows that azide coordinates directly to the metal without causing release of the metalcoordinated solvent molecule. Azide binding takes place without loss of protein ligands. The increased coordination number of the metal ion opens the His-Mn-His angle and the bent azide molecule across from the aspartate ligand. A similar arrangement was also observed for the Fe-superoxide dismutase from E. coli [72]. Note that in the case of the Cu,Zn-superoxide dismutase, cyanide, which acts as a competitive inhibitor of the enzyme, binds directly to the active site copper atom and replaces the bound water molecule. The coordination geometry of Cu(II) is partly altered with respect Met. Ions Life Sci. 2009, 6, 363–393


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to the uninhibited enzyme but its coordination number has not been increased [73,74]).

3.2.2. Manganese Catalase The Mn-catalases were originally identified on the basis of their insensitivity towards azide and cyanide. However, kinetic studies have subsequently shown that these enzymes are inhibited by azide and other anions, albeit at much higher concentrations than required for inhibition of heme catalases [59]. In the case of azide, the inhibition is competitive, suggesting that azide and peroxide bind at the same metal site. EPR spectroscopy first revealed that Mn catalase contained a binuclear Mn center, and this technique has been applied extensively to characterize the electronic structure of three of the four known oxidation states (see Table III.B in [70]). The distance between the Mn(II) varied between 3.31 A˚ (F) and B3.7 A˚ (unliganded, pH 7) for a variety of anions in a systematic way with size of the anion, consistent with binding to both Mn ions at a m-bridging position. Anions, with two or more lone pairs of electrons per atom, exhibited evidence of binding to the bridging site in the Mn(II),Mn(II) oxidation state and of simultaneously inhibiting the catalase activity. Note that cyanide showed a special behavior in that it did not inhibit catalase activity of the unliganded enzyme and actually reversed inhibition caused by the inhibiting anions. HCN, most likely, was the active form involved in reactivation of catalase activity. Cyanide appears to bind to a terminal (non-bridging) site on one or both Mn(II) ions, as seen by the increase in the intermanganese distance. This terminal ligation preference has been ascribed to the linear sp-hybridized electronic structure of cyanide. Apparently, bridging anions exchanged more slowly than terminally ligated anions and thus inhibited effectively. The stimulation of activity by cyanide suggests that bridging anions can be displaced to a labile terminal coordination site on one Mn ion.

4. NON-HEME IRON ENZYMES 4.1. Non-Heme Iron Enzymes. Structures and Functions Non-heme iron enzymes perform a wide range of important biological functions involving dioxygen in parallel to those of heme enzymes [75]. Oxygen-activating enzymes with mononuclear non-heme iron active sites participate in many metabolically important reactions that have environmental, pharmaceutical, and medical significance. For example, catechol Met. Ions Life Sci. 2009, 6, 363–393

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dioxygenases and Rieske dioxygenases are involved in the degradation of aromatic molecules in the environment. Lipoxygenases oxidize unsaturated fatty acids into precursors of leukotrienes and lipoxins and are potential targets for antiinflammatory drugs. Enzymes like isopenicillin N synthase and deacetoxycephalosporin C synthase, an a-keto acid-dependent enzyme, are important in the biosynthesis of antibiotics such as penicillin and cephalosporin [76]. Most mononuclear non-heme iron enzymes contain iron ligated by oxygen and/or nitrogen ligands. Usually, these enzymes promote dioxygen activation, resulting in the formation of highly reactive ironIV V peroxo (FeIII-OOH, FeIII -O 2 ) or iron-oxo (Fe ¼O or Fe ¼O) oxidation catalysts. The flexible coordination environment of enzymes containing 2-His-1-carboxylate (N2O)-ligated iron leaves room for substrate as well as dioxygen activation to occur at the metal site [77]. There also exists a class of iron-containing metalloenzymes utilizing an oxygen-bridged binuclear non-heme iron cluster, and some members of this class have been extensively characterized. The first member of the oxygenactivating binuclear iron class to be recognized, Fe-containing ribonucleotide reductase, is not an oxygenase during its primary catalyzed reaction, which is the reduction of ribonucleotides at the active site on its large subunit (R1). However, it does activate oxygen at a secondary active site on its small subunit (R2) in conjunction with the generation of a stable tyrosyl radical that is essential to the overall mechanism. The first true oxygenase of this class to be described was methane monooxygenase. The studies of this enzyme made it possible for the first time to compare the proposed oxygenactivating mechanism of P450 with that of a structurally dissimilar metallooxygenase. Recently, several other enzymes which are apparently similar to MMO, have been described in the literature [78].

4.1.1. Enzymes Carrying the 2-His-1-Carboxylate Facial Triad Structural Motif and Variants Great progress has been made in recent years toward our understanding of mononuclear non-heme Fe(II) enzymes thanks to a rapidly increasing number of crystal structures available for this class of enzymes. They activate dioxygen with a common structural motif, the iron(II) center in these enzymes is invariably coordinated by three protein residues, two histidines and one aspartate or glutamate, constituting one face of an octahedron, a recurring motif referred to as the 2-His-1-carboxylate facial triad (Figure 4) [79]. The 2-His-1-carboxylate facial triad serves as an excellent monoanionic three-pronged platform for binding divalent metal ions. The three remaining sites on the opposite face of the octahedron are consequently available for exogenous ligands. In the as-isolated enzymes, these sites are usually Met. Ions Life Sci. 2009, 6, 363–393


Figure 4.

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Structure of the 2-His-1-carboxylate facial triad motif [75].

occupied by solvent molecules but can accommodate both substrate (or co-substrate) and O2 in later steps of the catalytic cycle. Despite the many different transformations catalyzed, a general mechanistic pattern at the iron(II) center has emerged from spectroscopic and crystallographic studies of the various enzymes in this family. The Fe(II) center is six-coordinate at the start of the catalytic cycle and relatively unreactive toward O2. Subsequent substrate and/or cofactor binding to the active site makes the metal center five-coordinate and increases its affinity for O2. O2 binding then initiates the oxidative mechanism specific for each subclass. Thus, the metal center binds O2 only when substrate and cofactor(s) are present at the active site, thereby promoting strong coupling between the reduction of O2 and the oxidation of substrate. This structural motif thus allows the metal center to activate both substrate and O2 and bring them into close proximity for subsequent reaction, thereby accounting in large part for its versatility [80–82]. The application of the 2-His + Asp/ Glu Fe(II) binding motif documents how nature can catalyze a remarkable variety of oxygen activation chemistries with one functional structural building block. The organization of the second sphere environment can also be tuned by the strategic placement of acid/base catalysts, hydrogen bonding partners, non-bonding substrates, electron-supplying metal clusters, and cofactors. However, by controlling the structural elements of the Fe(II) center and its local environment, the point at which the O-O bond breaks during the reaction cycle can be regulated [83]. Relatively recently, a new class of cysteinate sulfur-ligated non-heme iron enzymes emerged which includes nitrile hydratases, superoxide reductases, and peptide deformylases. Superoxide reductase shares the N2X triad (X ¼ O, S) seen with the majority of non-heme iron enzymes, with a cysteinate (Figure 4) replacing the more common carboxylate residue, and two histidines replacing two of the waters. Nitrile hydratase and superoxide reductase have in common an active site containing a cysteinate sulfur trans to the substrate binding site. Two additional cysteinates ligate the iron of nitrile hydratase, completing one of the faces of an octahedron, with the Met. Ions Life Sci. 2009, 6, 363–393

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remaining coordination sites occupied by two peptide amide nitrogens and either a hydroxide or NO [77].

4.1.2. Protocatechuate 3,4-Dioxygenase Many soil bacteria express dioxygenases that are involved in the oxidative aromatic ring cleavage of catechol and its derivatives such as 3,4dihydroxybenzoate. These are important enzymes in the aerobic biodegradation pathways that allow these organisms to derive their carbon and energy from aromatic hydrocarbons. Depending on the position of the cleaved double bond relative to the hydroxyl groups, catechol dioxygenases can be split into two families: the intradiol-cleaving catechol dioxygenases, which cleave the carbon-carbon bond of the enediol moiety, and the extradiol-cleaving catechol dioxygenases, which cleave adjacent to the enediol. Although these enzymes share similar substrates, the intradiol- and extradiol-cleaving enzymes exhibit near exclusivity in their oxidative cleavage products, suggesting that there are two different mechanisms for cleavage. Furthermore, intradiol-cleaving catechol dioxygenases use an [FeIII(His)2(Tyr)2] active site, while extradiol-cleaving catechol dioxygenases contain a [MII(His)2(Asp/Glu)] active site, typically iron(II) but manganese(II) in a few cases [79]. Within this class of non-heme iron enzymes, perhaps the most extensively investigated is protocatechuate (3,4-dihydroxybenzoate) 3,4-dioxygenase. Crystallographic information is available for several complexes of the enzyme, including the as-isolated state, several enzyme-substrate complexes, and many complexes with inhibitors. The crystal structure of as-isolated protocatechuate (3,4-dihydroxybenzoate) 3,4-dioxygenase reveals a trigonal bipyramidal iron center, with four endogenous protein ligands (His460, His462, Tyr408, and Tyr447). The 5th coordination position, located in the trigonal plane, is occupied by a solvent-derived ligand [75,79].

4.1.3. The Non-Heme Iron Center of the Oxygen-Evolving Complex of Photosystem II A common property of the photosynthetic bacterial center and that of photosystem II of green plants is the presence of a non-heme iron center located between the quinones QA and QB. A number of spectroscopic similarities and sequence homologies suggest that in PSII-OEC, as in bacteria, the linear quinone-non heme iron-quinone arrangement spanning approximately 18 A˚ is conserved. Where the analogy deviates significantly is in the coordination and the properties of the iron. While in both systems Met. Ions Life Sci. 2009, 6, 363–393


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the metal is coordinated by four histidine residues, there are important differences in the 5th and 6th coordination positions. In bacteria, these positions are occupied by a bidentate glutamate residue [84], whereas in plants at least one of these positions is occupied by bicarbonate [85]. A number of carboxylate anions, or NO, can bind to the PSII-OEC non-heme iron center in competition with bicarbonate. In addition, the non-heme iron has been shown, unlike in its the bacterial reaction center homologue, to undergo redox changes between Fe(II) and Fe(III) [86]. Usually, in purple bacteria, as well as in algae and higher plants, the non-heme iron site appears to be in the reduced high-spin Fe(II) state, but can be oxidized to the high-spin Fe(III) state [87,88].

4.2. Non-Heme Iron Enzymes and Their Interaction with Cyanide 4.2.1. Enzymes Carrying the 2-His-1-Carboxylate Facial Triad Structural Motif and Variants Superoxide reductases remove potentially toxic superoxide anions from anaerobic organisms without forming O2 as a side product (reaction 10). In contrast, the better known superoxide dismutases produce one equivalent 2Hþ þ e þ O 2 ! H2 O2

ð10Þ

of O2 for every two molecules of O 2 (reaction 8). In the mechanism by which superoxide reductase presumably reduces O 2 , superoxide is proposed to bind to the reduced Fe(II) state of the enzyme at diffusion controlled rates (4109 M1 s1). The transfer of an electron from the Fe(II) center metal ion to the bound substrate via an innersphere pathway is then proposed to afford an Fe(III)-peroxide intermediate. Consistent with an innersphere pathway, exogenous ligands including azide, nitric oxide, and cyanide have been shown to bind to the iron site of superoxide reductase [89]. EPR and MCD studies provided evidence of azide, ferrocyanide, hydroxide, and cyanide binding via displacement of the glutamate ligand. For the first three ligands, ligand binding occurred with retention of the high-spin (S ¼ 5/2) ground state, whereas cyanide binding resulted in a low-spin (S ¼ 1/2) species, with g-values at 2.29, 2.25, 1.94. The ability to bind exogenous ligands to both the Fe(III) and the Fe(II) active sites is consistent with an innersphere mechanism for superoxide reduction. Most likely, superoxide binds at the vacant coordination site of the reduced enzyme coupled with electron transfer from iron to superoxide. Met. Ions Life Sci. 2009, 6, 363–393

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The cysteine ligand, which is in trans position to the superoxide binding site, would play a crucial role in pushing electron density on to the iron in order to promote Fe(II)-to-superoxide electron transfer, product dissociation from the Fe(III)-(hydro)peroxo intermediate, or both. Superoxide reductase is inhibited by cyanide but not by azide. A possible mechanism for cyanide inhibition would involve cyanide coordinating to the open coordination site of the catalytically active Fe(II) site. This would make the Fe site inaccessible to O 2 . However, this does not explain why superoxide reductase is not inhibited by azide. Another possibility is that CN inhibition involves the oxidized Fe(III) state. Experiments with model complexes showed that the redox properties of these complexes were dramatically altered by cyanide in comparison to azide. Presumably, cyanide prevents the enzyme from turning over by preventing the reduced, catalytically active Fe(II) state from being regenerated. Note that cyanide inhibits Fe-superoxide dismutase in a similar manner, by preventing the Fe(II) state from being regenerated [77,90].

4.2.2. Protocatechuate 3,4-Dioxygenase Protocatechuate 3,4-dioxygenase utilizes a Fe(III) center to catalyze the aromatic ring cleavage of 3,4-dihydroxybenzoate by incorporation of both atoms of dioxygen to yield b-carboxy-cis,cis-muconate. The crystal structures of several complexes of this enzyme with its substrate and with heterocyclic substrate analogs have been recently determined [91]. Complexation of the active site Fe(III) led to a dissociation of the endogenous axial tyrosinate ligand (Tyr447). After its release, Tyr447 becomes stabilized by hydrogen bonding and forms the top of a small cavity adjacent to the C3-C4 bond of the substrate. The equatorial Fe(III) coordination site within this cavity is unoccupied in the anaerobic enzyme-substrate complex but coordinates a solvent molecule in the complexes with the substrate analogs. In addition, ternary complexes with cyanide bound could be obtained. This shows that an O2 analogue can occupy the cavity and suggests that electrophilic O2 attack on the substrate 3,4-dihydroxy-benzoate is initiated from this site. Both the dissociation of the endogenous Tyr447 and the expansion of the iron coordination sphere are novel features of the 3,4protocatechuate 3,4-dioxygenase-substrate complex which appear to play essential roles in the activation of substrate for O2 attack.

4.2.3. The Non-Heme Iron Center of the Oxygen-Evolving Complex of Photosystem II Recently, it was shown that cyanide binds at the non-heme iron site of the PSII-OEC complex [92]. At lower pH, cyanide competed with NO for Met. Ions Life Sci. 2009, 6, 363–393


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binding to the iron (approximately 10 mM CN). At higher concentration of 21 EPR cyanide (approximately 50–80 mM), cyanide modified the Q A Fe signal at g ¼ 1.82–1.9 to a new form (g ¼ 1.98). It was assumed that at least two cyanide anions could bind to this iron site. More recently, in the 21 presence of NaCN (30–300 mM), at pH 6.5, the reduced state Q A Fe , produced by illumination at r200 K, or by reduction in the dark with sodium dithionite, was characterized by a g ¼ 1.98 EPR signal. This signal decayed with increasing pH above 6.5 and was almost absent at pH 8.1 and NaCN concentrations above 300 mM. Complementary to the disappearance of the g ¼ 1.98 EPR signal with increasing pH or incubation time, a new EPR signal developed which could be assigned to the uncoupled semiquinone Q A . These high pH, high cyanide concentration effects are accompanied by the conversion of the characteristic Fe21 (S ¼ 2) Mo¨ssbauer doublet to a new one with parameters characteristic of an Fe21 (S ¼ 0) state which explains the loss of the magnetic coupling of Q A with the iron center. A progressive binding of two or even three cyanide anions to the non-heme Fe21 site is proposed to explain the spectroscopic data [88]. The conversion of the iron to a diamagnetic state by cyanide treatment should help to obtain further valuable structural and functional informations on the Q A Fe21 site as important component of the PSII-OEC.

5. ZINC ENZYMES 5.1. Zinc in Biology. Structures and Functions Zinc is essential for growth and development in all forms of life. It is found in more than 300 enzymes, where it plays both a catalytic and a structural role. It is the only metal to have representatives in each of the six fundamental classes of enzymes: (i) oxidoreductases like alcohol dehydrogenase and superoxide dismutase, (ii) transferases like RNA polymerase and aspartate transcarbamoylase, (iii) hydrolases like carboxypeptidase A and thermolysin, (iv) lyases like carbonic anhydrase and fructose-1,6-bisphosphate aldolase, (v) isomerases like phosphomannose isomerase, and (vi) ligases like pyruvate carboxylase and aminoacyl-tRNA synthases [10,93]. There are growing numbers of nucleic acid binding proteins depending on Zn21, indicating that Zn21 is also widely involved in the regulation of the transcription and translation of the genetic message. Zn21 ion is redox inactive, as a consequence of its filled 3d10 configuration it does not have d-d transitions, and therefore no absorption in the visible region. Zinc is able to adopt a highly flexible coordination geometry, however, in most zinc proteins there is a strong preference for tetrahedral Met. Ions Life Sci. 2009, 6, 363–393

384

SOSA-TORRES and KRONECK X, Y, Z X

H Zn

Y

O H

Z

-O C 2

C(H) H3N+

-O C 2

H N CH2

N, O, S donors of His, Asp, Glu & Cys residues

O C(H)

N

CH2

OH

H3N+

His

-O C 2

C

Asp

-O

2C

O C(H)

CH2 SH

C(H)

CH2

CH2

C OH

H3N+

H3N+ Csy

Figure 5.

Glu

Common structural features of zinc enzymes [94].

coordination, frequently slightly distorted, which enhances both the Lewis acidity of the metal center and the acidity of a coordinated water molecule. Since Zn is of borderline hardness, it will bind oxygen (Asp, Glu, H2O), nitrogen (His) and sulfur (Cys) ligands (Figure 5).

5.1.1. Zinc Carbonic Anhydrase Carbonic anhydrase has played the most pivotal role in the development of enzymology [95–97]. It was the first enzyme recognized to contain a mononuclear zinc site, and is one of the most efficient enzymes known. It also has widespread occurrence in prokaryotes and has therefore been classified as an ‘‘ancient’’ enzyme [98]. The essential physiological function of the enzyme is to catalyze the hydration of carbon dioxide, and it thus plays an important role in respiration and intracellular CO2 =HCO 3 equilibration (equation 11). þ CO2 þ H2 O Ð HCO 3 þH

ð11Þ

These enzymes are very efficient catalysts for the reversible hydration of carbon dioxide to bicarbonate, but at least the a-class enzyme possesses a Met. Ions Life Sci. 2009, 6, 363–393


385

high versatility, being able to catalyze different other hydrolytic processes, such as (i) the hydration of cyanate to carbamic acid, (ii) the hydration of cyanamide to urea, (iii) the aldehyde hydration to gem-diols, (iv) the hydrolysis of carboxylic or sulfonic acid esters, and (v) other less investigated hydrolytic processes, such as hydrolysis of halogeno derivatives, arylsulfonyl halides, and other hydrolyzable substrates. It is not known whether reactions catalyzed by carbonic anhydrases other than the hydration of CO2 or dehydration of HCO 3 may have physiological relevance in organisms where these enzymes are present. The zinc ion is located at the bottom of a conical cavity (ca. 15 A˚ deep) and is coordinated to the protein by three histidine residues and a water molecule (or hydroxide ion, depending upon pH) forming a tetrahedral site (Figure 6). Recent crystallographic studies on the carbonic anhydrase from Methanosarcina thermophila (prototype of the g-class) revealed that the active site of this enzyme contained additional metal-bound water ligands, so the overall coordination geometry was trigonal bipyramidal [99]. The mechanism of action of carbonic anhydrase comprises three steps: (i) deprotonation of the coordinated water with a pKa E 7 to give [(His)3ZnOH]1, (ii) nucleophilic attack of the zinc-bound hydroxide at CO2 to give 1 the HCO 3 intermediate [(His)3Zn-OCO2H] , and (iii) displacement of the HCO3 anion by H2O to complete the catalytic cycle. Details concerned with the nature of the HCO 3 intermediate and how it is displaced from the metal center are still not fully understood [100,101]. To date, it is well established that carbonic anhydrase is widely distributed among phylogenetically and physiologically diverse prokaryotes, indicating a far greater role for this enzyme in nature than previously recognized. The comparison of sequences and crystal structures of the mammalian and plant enzymes demonstrates that they evolved independently and have

Glu-106 O N

His-94

H

Thr-199 O

N

N

H N

His-96

O

Zn

O H

His-119

N N

Figure 6.

Schematic drawing of the active site of carbonic anhydrase (a-class) [94]. Met. Ions Life Sci. 2009, 6, 363–393

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been designated the a-class and b-class, respectively. A third class, the g-class carbonic anhydrase, was reported in 1994, with a typical representative isolated from the methanogenic archaeon Methanosarcina thermophila [102].

5.1.2. Zinc Hydrolases Much of the importance of zinc enzymes derives from their peptidase and amidase activity involving the cleavage of RC(O)-NH(R 0 ) amide bonds. For example, with respect to peptidase activity, zinc enzymes include both endopeptidases (cleaving peptides or proteins at positions within the chain) and exopeptidases (cleaving a terminal amino acid from the chain). Of the exopeptidases, zinc enzymes function as both carboxypeptidases (which remove a C-terminal amino acid) and aminopeptidases (which remove a N-terminal amino acid). Other examples of zinc enzymes that function by cleaving amide bonds are (i) b-lactamases that destroy b-lactams (such as penicillin) by hydrolyzing and cleaving the four-membered lactam ring and (ii) matrix metalloproteinases that degrade extracellular matrix components such as collagen. In addition to the cleavage of amide bonds, zinc enzymes play an important role in the cleavage of the P-OR bond in phosphates, [(RO)PO3]2 and [(RO)2PO2], as exemplified by their nuclease activity pertaining to the hydrolysis of DNA and RNA. Examples of enzymes that incorporate two zinc centers include metallob-lactamases, aminopeptidases such as bovine lens leucine aminopeptidase, and alkaline phosphatases (Figure 7). There are also enzymes known that incorporate three zinc centers, such as phospholipase C and nuclease P1. Note that the third zinc centers of phospholipase C and nuclease P1 are not directly associated with the binuclear zinc site [94].

Glu

Asp X H2 O

His Zn

His His

OH2

η2 − Asp

Zn

Zn Asp

His

O

His

H2 O

O

O His Zn η2 − Glu O H2

Lys

Asp Zn

Zn Asp

O

Asp

O O

Asp

X = His, Cys

Figure 7. Schematic structures of the active sites of dinuclear zinc enzymes: metallob-lactamases and aminopeptidases [94]. Met. Ions Life Sci. 2009, 6, 363–393


387

5.2. Zinc Enzymes and Their Interaction with Cyanide 5.2.1. Zinc Carbonic Anhydrase Inhibitors and activators of the zinc enzyme carbonic anhydrase have a large number of applications in therapy. Many types of such new derivatives have been reported recently, together with their potential applications as antiglaucoma, anticancer and antiosteoporosis agents or for the management of a variety of neurological disorders, among others. Carbonic anhydrases are inhibited primarily by two main classes of compounds: the metal complexing inorganic anions, such as cyanide, cyanate, thiocyanate, azide, hydrogen sulfide, and the unsubstituted sulfonamides. Sulfonamide inhibitors are useful as diuretics, or in the treatment and prevention of a variety of diseases. Inhibitors of both types directly bind to the metal ion, either by substituting the Zn-bound H2O/OH, or adding to the coordination sphere, leading thus to pentacoordinated zinc. In the structure of the iodide complex of human carbonic anhydrase, the inhibitor I replaced the fourth coordinated H2O/OH ligand (Zn-I bond 2.7 A˚), whereas AuðCNÞ 2 was bound in a different part of the active site cavity and not directly to the Zn21 ion. The nitrogen atom of AuðCNÞ 2 is within hydrogen-bonding distance of the zinc-bound H2O/OH group which shifts by about 0.4 A˚ away from the zinc ion in relation to its position in the native enzyme. It is proposed that binding of the inhibitor AuðCNÞ 2 leads to a conformational reorientation of the activity-linked group, due to hydrogenbond formation with the inhibitor, which in turn sterically hinders the binding of the substrate CO2 molecule in the active site [103]. In a related study on the crystal structure of human carbonic anhydrase inhibited by cyanide and cyanate, it could be shown that the inhibitors replaced a water which formed a hydrogen bond to a peptide nitrogen (Thr199) in the native structure. The coordination of the Zn ion was hereby left unaltered, i.e., Zn21 coordinated three histidines and the H2O/OH group in a tetrahedral fashion. The binding site of the two inhibitors proved to be identical to what earlier has been suggested to be the position of the substrate, CO2, when attacked by the Zn-bound hydroxyl ion. The peptide chain underwent no significant alterations upon binding of either inhibitor [104]. Finally, in a recent study, the enzyme from the archaeon Methanosarcina thermophila was exposed to a large number of anions including halides, 2 CN, HCO 3 , CO3 , NO2 , HS , and HSO3 . The best Zn-carbonic 2 anhydrase inhibitors were HS and OCN , whereas SCN, N 3 , CO3 , and HSO3 were weaker inhibitors. For the Co-substituted enzyme, the metal poisons were less effective, with cyanide possessing an inhibition constant of approximately 50 mM [105]. Met. Ions Life Sci. 2009, 6, 363–393

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5.2.2. Zinc Hydrolases In experiments performed close to six decades ago, the effect of both zinc and magnesium ions on the activity of cyanide-inhibited (2 mM cyanide) kidney alkaline phosphatase was investigated. In low concentrations, Zn21 reactivated the inhibited enzyme whereas Mg21 had little effect [106]. A soluble form of alkaline phosphatase purified from Walterinnesia aegyptia snake venom was recently purified and characterized. Zinc and cyanide ions, at concentrations of 15 mM and 10 mM, respectively, completely inhibited the activity of the phosphatase [107].

6. CONCLUSIONS The cyanide anion, CN, and its transition metal complexes, are important compounds with interesting properties which turn them into valuable tools for both chemists and biologists. Based on the early chemistry of Prussian Blue, new cyano metal complexes have been designed which exhibit fascinating magnetic and electronic properties. The potential role of HCN as a precursor in prebiotic chemistry has been supported by the discovery that the hydrolytic products of its polymers including amino acids, purines, and orotic acid, a biosynthetic precursor of uracil, with the rapid formation of adenine by aqueous polymerization of HCN one of the key events in these studies. The cyanide anion is usually toxic for most aerobic organisms because of its inhibitory effects on respiratory enzymes on the one hand, but as a substrate it also represents an important form of carbon and nitrogen for many microorganisms, fungi, and plants. Finally, the cyanide anion is an important constituent of important metaldependent biomolecules, such as the hydrogenases and the cobalt site in vitamin B12.

ACKNOWLEDGMENTS This work was supported by Deutscher Akademischer Austauschdienst and Universidad Nacional Autońoma de Me´xico (M.E.S.T.) and Deutsche Forschungsgemeinschaft (P.M.H.K).

ABBREVIATIONS ADP ATP

adenosine 5 0 -diphosphate adenosine 5 0 -triphosphate

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CD EPR EXAFS MCD MMO NHE NMR PSII QA, QB ROS Sal SOD WOC

389

circular dichroism electron paramagnetic resonance extended X-ray absorption fine structure magnetic circular dichroism methane monooxygenase normal hydrogen electrode nuclear magnetic resoance photosystem II quinone centers A and B reactive oxygen species monoanion of salicylaldehyde superoxide dismutase water-oxidizing complex

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11 The Reaction Mechanism of the Molybdenum Hydroxylase Xanthine Oxidoreductase: Evidence Against the Formation of Intermediates Having Metal-Carbon Bonds Russ Hille Department of Biochemistry, The University of California, Riverside CA 92521, USA

ABSTRACT 1. INTRODUCTION 2. ELECTRON-NUCLEAR DOUBLE RESONANCE STUDIES OF THE ‘‘VERY RAPID’’ SPECIES 2.1. ENDOR of the Intermediate Seen with Xanthine as Substrate 2.2. ENDOR of the Intermediate Seen with 2-Hydroxy-6methylpurine as Substrate 3. X-RAY CRYSTAL STRUCTURES RELEVANT TO THE REACTION MECHANISM 3.1. Alloxanthine-Complexed Xanthine Oxidoreductase 3.2. Xanthine Oxidoreductase in Complex with FYX-051 3.3. Xanthine Oxidoreductase Reacting with 2-Hydroxy-6methylpurine and Lumazine 3.4. Desulfo Xanthine Oxidoreductase in Complex with Xanthine 3.5. Substrate Orientation and the Basis of Enzyme Catalysis 3.5.1. Glu730 3.5.2. Glu232 3.5.3. Arg310 Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00395

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4. GENERAL CONCLUSIONS ACKNOWLEDGMENTS ABBREVIATIONS AND DEFINITIONS REFERENCES

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ABSTRACT: ENDOR spectra of the catalytically relevant ‘‘very rapid’’ Mo(V) species generated in the course of the reaction of xanthine oxidoreductase with substrate have been examined by two different groups. While the data themselves are virtually identical, the analysis has been variously interpreted as supporting or refuting the existence of a molybdenum-carbon bond in the signal-giving species. While the basis for this difference in interpretation has now been generally agreed upon – the Mo-C distance in the signal-giving species is now understood to be too long to represent a direct Mo-C bond – independent information concerning the structure of the signal-giving species is highly desirable. Recently, several X-ray crystal structures of catalytically relevant complexes of the enzyme with several substrates and inhibitors have been reported. Taken together, these structures strongly and unambiguously support the interpretation that the intermediate giving rise to the ‘‘very rapid’’ EPR signal, as well as the Mo(IV) intermediate that precedes it in the reaction mechanism, has product coordinated to the active site molybdenum via the catalytically introduced hydroxyl group in a simple ‘‘end-on’’ fashion, with no metal-carbon bond character to the complex. The manner in which product is bound and its orientation within the active site provide important clues as to the specific catalytic roles of active sites in accelerating the reaction rate. KEYWORDS: catalysis molybdenum hydroxylase reaction mechanism xanthine oxidoreductase

1. INTRODUCTION The enzyme xanthine oxidoreductase catalyzes the final two steps of purine metabolism in most organisms (including humans), oxidatively hydroxylating hypoxanthine to xanthine and xanthine on to uric acid. The enzyme is most frequently encountered as a dehydrogenase that utilizes NAD1 to remove the reducing equivalents taken from substrate in the course of hydroxylation, but is also found as an oxidase that utilizes O2 rather than NAD1 [1]. The enzyme from cow’s milk (typically isolated as the oxidase) was first purified by Dixon and Thurlow in 1924 [2] and since then has been the subject of intensive investigation using a variety of sophisticated physicochemical methods, as well as both steady-state and rapid reaction kinetic methods. It was the first system to be examined using freeze-quench methodologies, following the formation and decay of paramagnetic Mo(V) intermediates formed in the course of the reaction with xanthine by electron paramagnetic resonance spectroscopy (EPR) [3,4]. This work identified several discrete Mo(V) species, ultimately designated ‘‘very rapid’’, ‘‘rapid’’, and ‘‘slow’’ on the basis of the kinetics of their formation and decay in the Met. Ions Life Sci. 2009, 6, 395–416

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course of the reaction of enzyme with xanthine [5]. The ‘‘slow’’ EPR signal was soon shown to arise from an inactive form of the enzyme lacking a labile sulfur at the molybdenum center (see below) [6]. The ‘‘rapid’’ species was subsequently shown to arise from a complex of partially reduced enzyme (formed by prior turnover) with substrate [7] and as such represented a paramagnetic analog of the ES Michaelis complex rather than a bona fide catalytic intermediate. It was immediately recognized that the ‘‘very rapid’’ species likely represented an authentic catalytic intermediate, and beginning in the early 1980’s Bray and coworkers embarked on an elegant series of isotopic substitution experiments aimed at elucidating the structure of the signal-giving species, and by implication the reaction mechanism. Using 8-[2H]-xanthine, it was shown that the C8 proton of substrate was initially transferred to a strongly magnetically coupled site of the molybdenum center, but was rapidly lost from this site by exchange with solvent [8]. Using 8-[13C]xanthine, observation of a small superhyperfine coupling demonstrated that the purine nucleus was an integral component of the signal-giving species [9]. It had been shown that treatment of enzyme with cyanide removed a catalytically essential sulfur from the molybdenum center (as thiocyanate) in a manner that could be reversed by incubation of the enzyme with sulfide under appropriate conditions [10], and 33S incorporated into the enzyme using Na233S. The ‘‘very rapid’’species generated with enzyme thus labeled exhibited strong and anisotropic coupling to the 33S nucleus and it was concluded that the sulfur was likely present as a terminal Mo¼S group in the active site molybdenum center [11,12]. This conclusion was subsequently confirmed by X-ray absorption spectroscopy (XAS) [13,14], which established the sulfur to be present at a distance of 2.15 A˚, consistent with a Mo¼S formulation. The XAS analysis also identified a Mo¼O at 1.68 A˚, consistent with a terminal Mo¼O group. Using EPR, a detailed analysis of the hyperfine coupling of 95Mo- and 97Mo-labeled enzyme (which constituted the initial evidence that molybdenum was an integral component of the enzyme active site [15]) was also interpreted in the context of a MoOS core [16]. In other isotope-labeling work, enzyme turnover in 17O-labeled water yielded a ‘‘very rapid’’ EPR signal exhibiting strong and isotropic coupling [17]. Presciently, it was concluded that this strongly coupled oxygen was not likely to be the Mo¼O group identified by XAS, but rather the bridging oxygen of a Mo-OR moiety that represented hydroxylated product coordinated to the molybdenum via the catalytically introduced oxygen atom (itself ultimately derived from solvent, as had been established previously [18]). Subsequent electron-nuclear double resonance (ENDOR) work established that apart from the Mo-OR oxygen, no other oxygen in the metal center, including the Mo¼O group, exchanged rapidly with solvent under turnover conditions [19]. Met. Ions Life Sci. 2009, 6, 395–416

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The basic elements of the active site molybdenum center established by EPR and XAS studies have been confirmed as crystal structures have become available, first of the closely related aldehyde oxidoreductase from Desulfovibrio gigas [20,21] and subsequently by the xanthine oxidoreductases from Bos taurus [22] and Rhodobacter capsulatus [23]. These structures have established the molybdenum coordination sphere of oxidized enzyme as LMoOS(OH) in a distorted square-pyramidal geometry (Figure 1). L here represents a pyranopterin cofactor coordinated to the molybdenum via an enedithiolate side chain, as shown. (Although this organic component of the metal center is frequently referred to as molybdopterin, this terminology is confusing on at least two counts: first, the term refers to the organic component alone, excluding the metal connoted by the prefix; and second, the same cofactor is found in tungsten-containing enzymes.) The Mo¼S group was originally assigned in the apical position of the molybdenum coordination sphere, which was surprising in light of the known structures of simple MoO-containing inorganic compounds. Subsequent work with the bovine enzyme [24] and related enzymes such as carbon monoxide dehydrogenase [25] and quinoline-2-oxidoreductase [26] has clearly established that it is the Mo¼O rather than Mo¼S that occupies the apical position. This assignment is consistent with the considerable literature on oxomolybdenum complexes [27,28]. In the protein, the Mo-OH ligand points into the solvent access channel toward the substrate binding site, a point of mechanistic significance as discussed below. With the active site of the oxidized enzyme established, other mechanistic information could next be incorporated to give a comprehensive picture of

S

S O Mo S OH

O H H2N

N N

H N

S

N H

O

Mo S OPi

Figure 1. Coordination geometry of the active site molybdenum center of XOR (top) and of the pyranopterin cofactor common to both molybdenum- and tungstencontaining enzymes (bottom). Met. Ions Life Sci. 2009, 6, 395–416


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the catalytic sequence. Most importantly, while it had long been known that the oxygen atom incorporated into product is ultimately derived from solvent [19], single-turnover experiments demonstrated that it is a catalytically labile site on the enzyme that is the proximal oxygen atom donor in the course of reaction, regenerated by oxygen from solvent at the completion of reaction [29]. Although originally thought to be the Mo¼O identified by XAS, it was subsequently established that a single turnover in 17O-labeled water incorporates 17O into a strongly and anisotropically coupled site in the molybdenum coordination sphere [30]. By analogy to EPR studies of isotopically labeled model compounds [31,32], it could be concluded that it is the Mo-OH rather than Mo¼O that represents the catalytically labile oxygen. This interpretation was subsequently substantiated when the reported crystal structures showed the Mo-OH ligand pointing directly at the substrate binding site in the protein, as discussed above. Other mechanistic information comes from the pH dependence of the reaction of enzyme with xanthine, where it has been shown that both the steady-state parameter kcat/Km and rapid reaction parameter kred/Kd (each reflecting the reaction of free enzyme with free substrate in the low-[S] regime), exhibit a bell-shaped pH dependence with pKa values of 6.6 and 7.4 from the acid and alkaline limbs of the curve, respectively [33]. The latter value agrees well with that for the ionization of neutral xanthine to the monoanion; the former pKa has been assigned to a universally conserved glutamate residue in the active site that is thought to act as an active site base [21,34]. Finally, use has been made of the slow substrate 2-hydroxy-6methylpurine, which in contrast to xanthine forms copious amounts of the ‘‘very rapid’’ EPR signal (and on a tens of seconds rather than milliseconds time scale [35]), to understand the kinetics of the reaction. Single-turnover experiments with this substrate have demonstrated that the reaction proceeds in three kinetic phases: the formation of a species having a difference maximum relative to oxidized enzyme at 470 nm, decay of this first species to one having a difference maximum at 540 nm, and finally the decay of this species back to oxidized enzyme (the experiment having been performed under aerobic conditions) [36]. The formation and decay of the 540 nmabsorbing species correlates well with formation and decay of the ‘‘very rapid’’ EPR signal, demonstrating that formation of the 540 nm-absorbing species is an oxidative event: an electron is lost from the enzyme in forming this species from the 470 nm-absorbing species that precedes it [36]. It can thus be concluded that the 470 nm-absorbing species represents a Mo(IV)product complex, which subsequently decays to the Mo(V)-product species giving rise to the ‘‘very rapid’’ EPR signal. Under the conditions used (pH 10 with 2-hydroxy-6-methylpurine as substrate) the ‘‘very rapid’’ species is essentially quantitatively generated. With other substrates and at moderate pH, however, much less of the signal-giving species is formed, presumably Met. Ions Life Sci. 2009, 6, 395–416

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due to dissociation of product from the Mo(IV)-product species prior to oxidation to the Mo(V) state.

2. ELECTRON-NUCLEAR DOUBLE RESONANCE STUDIES OF THE ‘‘VERY RAPID’’ SPECIES 2.1. ENDOR of the Intermediate Seen with Xanthine as Substrate Even though the ‘‘very rapid’’ species is not always generated to a significant degree under many (even most) experimental conditions, it remains extremely valuable as a paramagnetic reporter on the nature of the Mo-product interaction in the obligatory Mo(IV) species that precedes it in the catalytic sequence. To further investigate the manner in which product was bound to the active site molybdenum in the signal-giving species, Howes et al. examined the 13C coupling in the ‘‘very rapid’’ species generated with 8-[13C]-xanthine by ENDOR spectroscopy [19]. High quality data yielded a hyperfine tensor A ¼ [11.1, 7.6, 7.6] MHz, with the A tensor rotated from the z-axis of the g-tensor by 251. From the relationship A77+2A> ¼ 3 Aiso, the experimental A tensor yielded an Aiso of 8.8 MHz due to the small amount of spin density resident on C8 and an anisotropic term of the form T ¼ [2 T, -T, -T] due to the through-space interaction of the magnetic moments of the unpaired electron and the I ¼ 1/2 nucleus of the 13C nucleus, with T ¼ 1.17 MHz (reported as Aaniso¼3.5 MHz). This in turn was used to estimate a Mo-C distance of 2.2 A˚ in the signal-giving species. This distance estimate was recognized to be dependent on the hybridization of the C8 carbon and was taken as an upper limit; a more realistic distance of B2.1 A˚ was subsequently estimated assuming that sp hybridization yielded the longest possible distance (this in turn was based on the assumption that an sp-hybridized orbital was less spherically symmetric than an sp3-hybridized one). On the basis of the conclusion that the C8 of bound product was likely to be within bonding distance of the molybdenum in the ‘‘very rapid’’ species, a reaction mechanism was proposed in which the C8-H bond of substrate inserts across the Mo¼S bond of oxidized enzyme to yield an intermediate in which the C8 carbon directly bonds to the active site molybdenum. A ‘‘buried water’’ (the authors specifically considered the possibility that it was coordinated to the molybdenum [17]) was then proposed to attack the Mo-C bond to give an intermediate in which the C8 carbonyl of the newly formed uric acid was coordinated to the now reduced molybdenum in a side-on Z2 fashion, in which the Met. Ions Life Sci. 2009, 6, 395–416


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N S

O VI

S

Mo S

O

S

N H

S

N

Mo

N N

VI

S H

SH

H2O

2 H+

S

O

N

H

Mo O SH

H

e-, H+

N N S

O VI

S

Mo S

OH +

S

N

O

N H

S

S

N H2O

N

H

V

VI

H

O

Mo O S

S +

H

e-

Mo O S

Figure 2. The reaction mechanism proposed by Bray and coworkers [17], on the basis of a short Mo-C distance in the species giving rise to the ‘‘very rapid’’ EPR signal. The reaction is proposed to begin with the insertion of the C8-H bond across the Mo¼S group of the molybdenum center to give the species shown with no reduction of the molybdenum itself.

p electrons of the carbonyl bond donate into the molybdenum dxy orbital (Figure 2). This bonding interaction was taken as a vestige of the direct Mo-C bond of the previous intermediate. The reaction was completed by sequential oxidation of the molybdenum and displacement of product by hydroxide from solvent. Precedent for insertion chemistry of the type proposed exists in the reactivity Mo¼S units within complexes such as the binuclear [MoOS(m-S)2MoO(Cp)]2, which are susceptible to insertion reactions with compounds such as carbon disulfide or dicarbomethoxyacetylene [37].

2.2. ENDOR of the Intermediate Seen with 2-Hydroxy-6methylpurine as Substrate The interpretation that the Mo-C8 distance in the ‘‘very rapid’’ species was sufficiently short to be interpreted as a direct Mo-C bond was controversial and prompted a second ENDOR study examining the ‘‘very rapid’’ signal generated using 8-[13C]-2-hydroxy-6-methylpurine as substrate [38]. This study was facilitated by the high levels of signal-giving species afforded by this substrate (and with the signal-giving species forming on a more convenient time scale than with xanthine), and yielded A ¼ [10.2, 7.0, 6.5] MHz. Met. Ions Life Sci. 2009, 6, 395–416

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While modestly different from the values obtained with xanthine as substrate, the value for T of 1.15 MHz derived with 2-hydroxy-6-methylpurine is essentially indistinguishable from the value of 1.17 MHz seen with xanthine, and yielded an identical apparent Mo-C distance of 2.2 A˚. These workers noted, as did Howes et al. [18], that T has an anisotropic local contribution arising from the finite spin density on C8 and that only the non-local (through-space or dipolar) contribution to T should be used in the distance calculation. Tloc is dependent on the degree of hybridization at C8, introducing some uncertainty into its estimation, but a table was constructed demonstrating that the Mo-C distance increased from 2.4 A˚ with sp hybridization to 2.7 A˚ with sp2, and 3.0 A˚ with sp3 hybridization. That the correction increased with the degree of p hybridization arises from the fact that a given sp orbital is more spherically symmetric than a given sp3 orbital, a point not appreciated in the earlier work. Assuming nominal sp2 hybridization and taking into account the finite spin density on the catalytically introduced oxygen, a value of 2.8 A˚ was estimated for the Mo-C distance [38]. This was deemed too long for a direct Mo-C bond, and was in fact in good agreement with the distance expected for product coordinated to the molybdenum via the catalytically introduced oxygen with a Mo-O-C bond angle of 1091. This geometry is depicted in Figure 3, which also illustrates that this geometry is also consistent with Mo-H distances of 3.2 and 6.1 A˚ for the N7-H and 6-methyl protons (averaged) of 2-hydroxy-6-methylpurine, as previously determined using pulsed EPR methods [39]. It is now generally accepted that the ENDOR data support a Mo-C distance too great to reflect formation of a direct Mo-C bond in the course of the hydroxylation reaction. As has been reviewed elsewhere [40–42], the catalytic sequence is instead envisaged as being initiated by abstraction of the Mo-OH proton by Glu1261 in the bovine structure

H

2.8 Å

N

N

Mo

O

O N

N H

3.2 Å

6.1 Å avg

C H

H H

Figure 3. A summary of the metric information for the ‘‘very rapid’’ species obtained with 2-hydroxy-6-methylpurine from ESEEM [38] and ENDOR [39] analysis. The Mo-O-C8 bond angle agrees well with that subsequently seen in the crystal structure of the intermediate [47]. Met. Ions Life Sci. 2009, 6, 395–416

MOLYBDENUM HYDROXYLASE XANTHINE OXIDOREDUCTASE S O VI S Mo S

N H

O H

O Glu1261

H N

N H

-

O

O N

O

H

S O IV S Mo SH O

N

N H

H N N O

S OV S Mo S O

O

H+, e-

H

N N H

H N N O

O

403

HO-, H+

P, e-

S O VI S Mo S OH

H

e-

P

e-

Figure 4. A reaction mechanism for xanthine oxidoreductase predicated on transfer of a Mo-OH moiety on C8 of substrate, as originally proposed by Wedd and coworkers [32]. The position of the Mo-OH in the active site, as subsequently seen in the crystal structure of the D. gigas aldehyde oxidoreductase [20], was appropriate for such group transfer, and it was suggested that an active site glutamate (Glu1261 in the structure of the bovine xanthine oxidoreductase) functioned as an active site base to deprotonate the Mo-OH and initiate catalysis.

(Glu730 in the R. capsulatus enzyme), followed by nucleophilic attack on the substrate site to be hydroxylated and concomitant hydride transfer from C8 of substrate to the Mo¼S group, as depicted in Figure 4. The initial intermediate can be formulated as LMo(IV)O(SH)OR, and in this first step of the reaction the C-H bond of substrate is broken, O-C bond of product is formed, and the molybdenum center formally reduced from the (VI) to (IV) oxidation state. This key intermediate may decay either by transfer of an electron to other redox-active centers in the enzyme (it possesses two [2Fe-2S] iron-sulfur clusters and FAD in addition to the molybdenum center) to give the ‘‘very rapid’’ species, or by displacement of bound product by hydroxide from solvent prior to electron transfer out of the molybdenum center, thereby circumventing formation of the EPR-active intermediate. Decay of the ‘‘very rapid’’ species, to the extent that it is formed, occurs by displacement of product by hydroxide and concomitant transfer of the second electron out of the molybdenum center. Such a mechanism, involving transfer of a Mo-OH ligand to substrate in forming product, was first suggested by Wedd and coworkers [32] on the basis of a comparison of 17O hyperfine coupling seen in the EPR of a series of structurally defined model compounds with that observed in various enzyme species. The disposition of the Mo-OH facing into the solvent access channel to the active site in the enzyme, as established by subsequent crystallographic work, is consistent with such a mechanism. The available data are thus fully consistent with a reaction mechanism that proceeds as indicated above and shown in Figure 4, with product coordinated to molybdenum in the initially formed EredP intermediate in a simple end-on fashion via the catalytically introduced oxygen. Met. Ions Life Sci. 2009, 6, 395–416

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3. X-RAY CRYSTAL STRUCTURES RELEVANT TO THE REACTION MECHANISM Over the past several years a number of X-ray crystal structures in addition to that of oxidized enzyme have appeared, including several that are of direct mechanistic significance specifically regarding the structure of the EredP intermediate. What follows is a discussion of these structures, then a consideration of the catalytic roles of specific active site residues in the context of our present understanding of substrate orientation within the active site.

3.1. Alloxanthine-Complexed Xanthine Oxidoreductase Allopurinol (4-hydroxypyrazolo[3,4-d]-pyrimidine) was developed in the late 1960’s as an inhibitor of xanthine oxidoreductase to treat hyperuricemia and as a tandem drug to potentiate the effectiveness of 6-mercaptopurine and related first-generation chemotherapeutic agents [43]. George Hitchings and Gertrude Elion shared the 1988 Nobel Prize in Physiology or Medicine (with Sir James Black, for unrelated work) for the successful development of this tandem drug therapy in the treatment of cancer. Allopurinol was found to be oxidatively hydroxylated to alloxanthine (4,6-dihydroxypyrazolo[3,4d]-pyrimidine) in a manner analogous to the conversion of hypoxanthine to xanthine, but with the product alloxanthine bound tightly to the reduced form of the molybdenum center to inhibit the enzyme [44]. Interest in the structure of the inhibitory complex was greatly stimulated by the discovery that partial oxidation of the Eredalloxanthine complex resulted in an EPR signal that was strongly reminiscent of that of the ‘‘very rapid’’ species, with unresolved superhyperfine structure due to coupling to a nitrogen nucleus [45,46]. It was immediately recognized that the likely structure of the signalgiving species involved direct coordination of the inhibitor to the molybdenum via the nitrogen at the position analogous to C8 in xanthine. (It was also inferred at the time that the ‘‘very rapid’’ species involved the product uric acid, coordinated via the catalytically introduced oxygen, an interpretation later abandoned in favor of a direct Mo-C bond, as discussed above). The crystal structure of alloxanthine-complexed enzyme was first reported for the xanthine dehydrogenase from Rhodobacter capsulatus [23], which has a very high degree of sequence and structural homology to the vertebrate enzymes. As illustrated in Figure 5, the structure shows the inhibitor bound in the active site between two phenylalanine residues (one interacting face-on, the other side-on with the aromatic inhibitor), and coordinated to the molybdenum via N2, just as predicted on the basis of the EPR work. Inhibitor is oriented in the active site such that its C6-carbonyl is directed Met. Ions Life Sci. 2009, 6, 395–416


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Figure 5. The structure of the reduced R. capsulatus xanthine dehydrogenase in complex with the tight-binding inhibitor alloxanthine [22]. The orientation of the pyrimidine ring of substrate is such that its C6 carbonyl (equivalent to the C2 carbonyl of xanthine) is oriented toward the conserved Arg310 in the active site.

toward Arg310. The structure of the alloxanthine complex defines the substrate binding site, with complexed product occupying the position otherwise occupied by the Mo-OH group in the coordination sphere of oxidized enzyme.

3.2. Xanthine Oxidoreductase in Complex with FYX-051 The next structure of mechanistic relevance is that of the bovine enzyme in complex with the inhibitor FYX-051, another pharmaceutically important inhibitor of the enzyme (marketed as Febuxostat). Like allopurinol, FYX051 is hydroxylated by enzyme [24], but unlike alloxanthine product remains tightly coordinated to the molybdenum center via the catalytically introduced hydroxyl group. In this sense, FYX-051 is a true mechanismbased inhibitor, as opposed to allopurinol which is simply a target-activated drug. As shown in Figure 6, the crystal structure of the reduced enzymeinhibitor complex shows the hydroxylated product coordinated to the molybdenum center with a Mo-O-C geometry closely resembling that predicted on the basis of the ENDOR/ESEEM data discussed above (Figure 3), with a Mo-O-C bond angle of B1101. Like alloxanthine, FYX-051 has a hydrophobic portion that inserts between the two active site phenylalanine residues. FYX-051 is a much larger molecule than is alloxanthine, however, and its therapeutic effectiveness appears to derive from a close structural complementarity to most (if not all) of the solvent access channel of the mammalian enzyme. Interestingly, FYX-051 does not inhibit the R. capsulatus enzyme, presumably because the solvent access channel of the bacterial enzyme is shaped differently than in the mammalian enzyme (a point suggesting that species-specificity may in at least some cases be engineered into mechanism-based inhibitors). Met. Ions Life Sci. 2009, 6, 395–416

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Figure 6. The crystal structure of bovine xanthine oxidoreductase in complex with the inhibitor FYX-051 [23]. The complex is of a particularly stable intermediate formed in the course of hydroxylation of the inhibitor, corresponding to EredP, in which the hydroxylated product is coordinated to the metal center via the catalytically introduced hydroxylate.

3.3. Xanthine Oxidoreductase Reacting with 2-Hydroxy-6methylpurine and Lumazine More recently, the crystal structure of the bovine enzyme during turnover with the slow substrate 2-hydroxy-6-methylpurine has been determined [47]. This substrate, which generates large amounts of the ‘‘very rapid’’ species as discussed above, reacts on a sufficiently slow time scale (tens of seconds) that it is possible to arrest the enzyme during catalysis simply by briefly soaking pre-grown crystals of the enzyme with substrate prior to freezing the crystal for data acquisition. As seen in Figure 7, product is found bound in the active site, inserted again between the two phenylalanine

Figure 7. The crystal structure of bovine xanthine oxidoreductase in complex with 2-hydroxy-6-methylpurine and lumazine, and of desulfo enzyme with xanthine. As with FYX-051, the first two complexes represent particularly stable catalytic intermediates in which the hydroxylated substrate coordinates the metal center via the catalytically introduced hydroxylate. The orientation of the pyrimidine subnucleus of 2-hydroxy-6-methylpurine (top) is comparable to that of alloxanthine in the reduced R. capsulatus protein, with the C2 carbonyl oriented toward Arg880. The orientation with lumazine, however, is inverted with the C2 carbonyl (equivalent ot C6 of xanthine) oriented toward the arginine (middle). For xanthine bound to the (in)active site of the desulfo enzyme the pyrimidine subnucleus of substrate oriented such that its C6 carbonyl is oriented toward Arg880 (bottom), analogous to the orientation seen with lumazine but opposite that seen with 2-hydroxy-6-methylpurine. Met. Ions Life Sci. 2009, 6, 395–416


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residues, and coordinated to the molybdenum via the catalytically introduced hydroxyl group in a manner directly analogous to the structure seen with the inhibitor FYX-051. The sole carbonyl of the pyrimidine subnucleus, at C2, is oriented toward Arg880 (Arg310 in the R. capsulatus enzyme), with the purine in an orientation analogous to that of alloxanthine seen in its complex with reduced enzyme. There is clear electron density bridging between the C8 carbon and molybdenum [47], and the structure is clearly that of the Eredproduct complex for this slow substrate. The structure provides compelling support for the interpretation above of the ENDOR data regarding the nature of product binding in the EredP complex. On the basis of a best estimate for the Mo-S distance in the equatorial plane (2.0 0.2 A˚), it would appear that the complex is that of the Mo(V)¼S species (rather than the Mo(IV)-SH species) that gives rise to the ‘‘very rapid’’ EPR signal, although this could not be definitively established given the well-known difficulties associated with Fourier transform truncation artifacts introduced into the electron density in the vicinity of atoms as heavy as molybdenum [48]. Interestingly, the structure described above is seen in only one of the two active sites of the homodimeric enzyme (of which one dimer is present in the asymmetric unit of the crystal). In the second site, the purine nucleus sits somewhat further back from the molybdenum center with no clear evidence of intervening electron density [47]. The structure is apparently that of the Michaelis complex of the enzyme, with substrate situated in the active site in an appropriate orientation for catalysis, but with no evidence of the catalytic sequence having been initiated. The implication is that in the crystallographically defined dimer, only one of the two subunits has progressed into the catalytic sequence. This suggests that, in the crystal if not in free solution, the enzyme may function in a reciprocating fashion, with first one then the other subunit functioning catalytically. Such half-sites reactivity for the enzyme has been suggested previously [49], but given the absence of any obvious structural basis for the implied subunit-subunit interaction (the two subunits in the crystal structure appear to have exactly the same protein folds) it remains to be determined whether the enzyme indeed functions in such a way. The structure of the same intermediate seen in the course of the reaction of the bovine enzyme with lumazine (2,4-dihydroxypteridine) has also been determined very recently (J. M. Pauff, H. Cao, and R. Hille, unpublished). Lumazine is the pterin analog to the purine xanthine, and has been shown to be hydroxylated by xanthine oxidoreductase to give violapterin (2,4,7-trihydroxypteridine) [50]. In the course of the reaction a long-lived intermediate absorbing at long wavelength is formed that has been shown to represent a catalytically competent EredP species [51]. As compared with 2-hydroxy-6-methylpurine, lumazine binds with its pyrimidine subnucleus Met. Ions Life Sci. 2009, 6, 395–416


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oriented such that the C2 carbonyl (equivalent to C6 in xanthine) points toward Arg880, i.e., inverted relative to the orientation seen with 2-hydroxy6-methylpurine, as shown in Figure 7. As discussed further below, this is also the orientation seen with xanthine bound to the protein and appears to represent the catalytically preferred orientation for substrate binding.

3.4. Desulfo Xanthine Oxidoreductase in Complex with Xanthine The final crystal structure of catalytic relevance is (ironically) that of the inactive desulfo form of the bovine enzyme in complex with xanthine (J. M. Pauff, H. Cao, and R. Hille, unpublished). As shown in Figure 7, the structure shows substrate bound at the active site with the C6 carbonyl oriented toward Arg880 (Figure 8); it is evident from the shape of the electron density that an inverted orientation similar to that seen with 2-hydroxy-6-methylpurine does not fit well into the observed electron H O S

Mo

N S

H

O N

N H

O-

S

N

H

O NH2

H2N

R880

H

O S S

Mo

S O

H

N

N

N H

O-

O N

H2N

H

NH2 R880

Figure 8. Proposed stabilization of negative charge accumulation on the C6 carbonyl of xanthine in the course of nucleophilic attack on substrate. Stabilization of negative charge on C2 in a substrate orientation inverted relative to that shown is expected to be less effective given the expected lower degree of negative charge distribution onto C2. Met. Ions Life Sci. 2009, 6, 395–416

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density. This orientation is inverted from that seen with 2-hydroxy-6methylpurine (where the C2 carbonyl is oriented toward the active site arginine) but analogous to the orientation seen with lumazine above. In an earlier kinetic study of a homologous series of purine substrates, these could be segregated into two distinct groups: those that were effective substrates of wild-type enzyme and which were profoundly affected by mutation of Arg310 (in the R. capsulatus enzyme), with kred in the reductive half-reaction reduced by approximately 104; and those that were poor substrates but which were not profoundly affected by the mutation [52]. It was suggested that the dichotomy in substrate reactivity was due to differences in the orientation of substrates in the active site. Effective substrates had a C6 carbonyl (or thio) group and were proposed to bind in the active site as seen with xanthine, with this group oriented toward Arg310 (Arg880 in the bovine sequence). This arginine appeared positioned to stabilize negative charge accumulation on the heterocycle in the course of nucleophilic attack by charge complementation at the C6 carbonyl oxygen, as shown in Figure 8. Substrates that were less effective (e.g., 2-hydroxy-6methylpurine) bound in the opposite orientation, with C2 oriented toward Arg310. Although not reacting as rapidly with wild-type enzyme, these poorer substrates were also much less sensitive to mutation of Arg310 to Met – in the case of 2,6-diaminopurine, the substrate is actually somewhat more reactive with the R310 M mutant than wild-type enzyme [52].

3.5. Substrate Orientation and the Basis of Enzyme Catalysis The above crystallographic information provides crucial information not only about the structures of the critical Eredproduct and ‘‘very rapid’’ intermediates in the catalytic sequence (and by inference the specific mechanism by which substrate is chemically converted to product) but also the roles of specific amino acid residues in accelerating reaction rate by establishing the orientation of substrate in the active site. We consider here three specific residues, Glu767, Glu1261, and Arg880 (Glu232, Glu730, and Arg310 in the numbering system of the R. capsulatus xanthine dehydrogenase, with which the site-directed mutagenesis studies to be discussed were performed). The role of each of these residues, depicted in Figure 9, in catalysis will now be considered.

3.5.1. Glu730 Glu730 is universally conserved in the purine- and aldehyde-hydroxylating molybdenum enzymes, and as indicated above on the basis of the crystal Met. Ions Life Sci. 2009, 6, 395–416


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Figure 9. Residues of the active site of xanthine oxidoreductase that interact with substrate or the molybdenum center of the active site.

structure of the first of these enzymes to be determined (that of the aldehyde oxidoreductase from D. gigas) is thought to act as a general base, deprotonating the Mo-OH of the molybdenum center to facilitate nucleophilic attack on substrate (be it a purine as illustrated in Figure 4, or an aldehyde) [21]. The protonation state of the ligand to the metal is important, and although the initial X-ray crystallographic work concluded that the ligand was a doubly protonated water molecule, it has subsequently been established using the more precise method of X-ray absorption spectroscopy (with the bovine enzyme) that it is a singly protonated Mo-OH [53], whose deprotonation leads to the much more nucleophilic Mo-O. Mutation of Glu730 profoundly affects catalysis: while the wild-type enzyme is fully reduced by 100 mM xanthine within 100 ms under anaerobic conditions at pH 7.8, the E730A mutant is not perceptibly reduced in the course of an overnight incubation with substrate at the same concentration [34]. The effect on the limiting rate of reduction is conservatively estimated to be a factor of 107, amounting to B10 kcal/mol of transition state stabilization contributed by Glu730. Interestingly, XAS analysis of the (bovine) enzyme at pH 10 suggests a shortening of the Mo-O bond consistent with deprotonation [53] and indeed much of the disparity in reactivity between wild-type and E730A mutant is lost in going from pH 7.4 to 10 (Ibdah and Hille, unpublished). The effect is approximately equally due to Met. Ions Life Sci. 2009, 6, 395–416

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loss of activity of wild-type enzyme as gain of activity of the mutant at high pH.

3.5.2. Glu232 Glu232 sits opposite Glu730, atop the active site as shown in Figure 9, and is strictly conserved only among enzymes that hydroxylate xanthine – it is typically absent in aldehyde-oxidizing enzymes. Mutation of this residue to an alanine results in a B10-fold decrease in the limiting rate of enzyme, kred, reduction at high [xanthine] and a similar B10-fold increase in Kd for xanthine – this residue thus contributes approximately 1.5 kcal/mol to transition state stabilization and a similar amount to substrate affinity [34]. In a computational analysis of tautomeric forms of substrate likely to be encountered in the course of catalysis [54], it was found that the tautomer with protons on nitrogens 1, 7, and 9 was significantly stabilized relative to the predominant tautomer in solution (with protons on nitrogens 1, 3, and 7) once nucleophilic attack has occurred (Figure 10). Glu232 was proposed to

H N

H N

H H H N

O N

H H N

H

O H N

H

O N

N

1H, 3H, 9H

H

N9 H 1H, 3H, 7H H

1H, 3H, 9H

O N1

N7 O

N

H

N H

O

H N

H N

1H, 3H, 7H

O

H

H

N

N H

O

N

H N

N H

O N

H

O N

H

O

23 kcal/mol

O N

N H

H N

H Mo

O H N

H

1 kcal/mol H

N

N

N Free Energy

O N

H

O H N3

Mo

H 1H, 7H, 9H

H N

O N

N O

N

N H

H N

O

H 1H, 7H, 9H

Mo

O

H N

N

N

3H, 7H, 9H

H

H

O

substrate

sp3 intermediate

Figure 10. Possible role of tautomerism in the course of the reaction of xanthine oxidoreductase. Left: the four tautomeric forms of neutral xanthine. Right: the relative stabilities of the three most stable tautomeric forms of xanthine in solution and as part of an EredP complex. Met. Ions Life Sci. 2009, 6, 395–416


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facilitate this tautomerization, accounting for the rate acceleration attributable to this residue. The orientation of substrate seen in the crystal structure of desulfo (bovine) xanthine oxidase in complex with xanthine is consistent with such a role. It has alternatively been proposed that this tautomerization is facilitated by Glu730 [55], and it would indeed be elegant were Glu730 to simultaneously deprotonate the Mo-OH and transfer the proton thus obtained to N9 of substrate once nucleophilic attack has been initiated. Such a role, however, is predicated on a substrate orientation in the active site opposite to that seen crystallographically and now seems unlikely.

3.5.3. Arg310 As indicated above, it is the orientation of substrate relative to Arg310 that is proposed to determine whether a given purine substrate will be an effective or ineffective substrate for enzyme, due to the ability of the arginine to stabilize negative charge accumulation on the C6 carbonyl oxygen in the course of nucleophilic attack. Although stabilization via the C2 carbonyl may occur to some degree when substrate is in the unproductive orientation, it is expected to be less effective than stabilization via the C6 carbonyl as evidenced in the Kekule´ structures shown in Figure 8, where it can be seen that it is more difficult to delocalize negative charge on the C2 carbonyl oxygen. Thus as with Glu232, substrate orientation dictates the role of Arg310 in catalysis.

4. GENERAL CONCLUSIONS Although the formation of a direct Mo-C bond in the course of the reaction of xanthine oxidoreductase was advocated on the basis of the analysis of the original ENDOR data obtained with the ‘‘very rapid’’ intermediate, subsequent analysis strongly suggests that the Mo-C distance is no shorter than 2.8 A˚ and that it is unlikely that the signal-giving species possesses a direct bond between metal and substrate carbon. Several subsequently determined X-ray crystal structures, including those of catalytic intermediates seen with FYX-051, 2-hydroxy-6-methylpurine and lumazine, support the latter ENDOR interpretation of substrate binding in a simple end-on fashion in the critical intermediate(s). The structures imply a reaction mechanism that is initiated by base-assisted nucleophilic attack on substrate after deprotonation of a Mo-OH group of the molybdenum center. Such a mechanism provides a ready explanation for the observed effects seen on mutagenesis in the active site of the enzyme. Met. Ions Life Sci. 2009, 6, 395–416

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ACKNOWLEDGMENTS Work in the author’s laboratory has been supported by the National Institutes of Health (GM 075036). The author wishes to acknowledge Mr. J. M. Pauff for preparation of the structural Figures 5, 6, 7, and 9.

ABBREVIATIONS AND DEFINITIONS allopurinol alloxanthine ENDOR EPR ESEEM FAD lumazine NAD1 violapterin XAS XOR

4-hydroxypyrazolo[3,4-d]-pyrimidine 4,6-dihydroxypyrazolo[3,4-d]-pyrimidine electron-nuclear double resonance electron paramagnetic resonance electron spin echo envelope modulation spectroscopy flavin adenine dinucleotide 2,4-dihydroxypteridine nicotinamide adenine dinuclotide 2,4,7-trihydroxypteridine X-ray absorption spectroscopy xanthine oxidoreductase

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12 Computational Studies of Bioorganometallic Enzymes and Cofactors Matthew D. Liptak, Katherine M. Van Heuvelen, and Thomas C. Brunold * Department of Chemistry, University of Wisconsin-Madison, Madison WI 53706, USA

ABSTRACT 1. INTRODUCTION 2. COMPUTATIONAL APPROACHES TO BIOORGANOMETALLIC CHEMISTRY 2.1. Overview 2.2. Popular Methods for Computing Geometric and Electronic Structures 2.3. Computation of Spectroscopic Observables 3. FORMATION AND CLEAVAGE OF THE CO–C BOND OF COBALAMIN IN ENZYME ACTIVE SITES 3.1. Overview 3.2. Co–C Bond Formation During Adenosylcobalamin Biosynthesis and Methylcobalamin Activation by Methionine Synthase 3.3. Co–C Bond Homolysis in Adenosylcobalamin-Dependent Enzymes 3.4. Co–C Bond Heterolysis in Methylcobalamin-Dependent Enzymes 4. ORGANOMETALLIC CHEMISTRY AND CATALYTIC CYCLE OF METHYL-COENZYME M REDUCTASE Metal Ions in Life Sciences, Volume 6 Edited by Astrid Sigel, Helmut Sigel, and Roland K. O. Sigel r Royal Society of Chemistry 2009 Published by the Royal Society of Chemistry, www.rsc.org DOI: 10.1039/9781847559159-00417

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4.1. Overview 435 437 4.2. Properties of the Free F430 Cofactor 4.3. Ni-Alkyl Species of Methyl-Coenzyme M Reductase: Computational Evaluation of Viable Reaction Mechanisms 439 4.3.1. Ni–Methyl Pathway 439 4.3.2. Methyl Radical Pathway 441 442 4.3.3. Protonation of Ni(I)MCRRed1 5. GEOMETRIC AND ELECTRONIC STRUCTURES OF THE CARBON MONOXIDE DEHYDROGENASE/ACETYLCOENZYME A SYNTHASE ACTIVE SITE CLUSTERS 442 5.1. Overview 442 5.2. Mechanism of Carbon Monoxide Oxidation/Carbon Dioxide Reduction by Carbon Monoxide Dehydrogenase 444 5.3. Computational Insights into the Catalytic Cycle of AcetylCoenzyme A Synthase 445 6. MAGNETIC PROPERTIES OF THE ACTIVE SITE CLUSTER OF IRON-ONLY HYDROGENASES 447 6.1. Overview 447 6.2. Structure and Oxidation States of the Active Site Cluster of [FeFe] Hydrogenases 447 6.3. Calculation of the Magnetic Properties of the H-Cluster 449 7. CONCLUDING REMARKS AND FUTURE DIRECTIONS 450 ACKNOWLEDGMENTS 452 ABBREVIATIONS 452 REFERENCES 454 ABSTRACT: Because of their complex geometric and electronic structures, the active sites and cofactors of bioorganometallic enzymes, which are characterized by their metal–carbon bonds, pose a major challenge for computational chemists. However, recent progress in computer technology and theoretical chemistry, along with insights gained from mechanistic, spectroscopic, and X-ray crystallographic studies, have established an excellent foundation for the successful completion of computational studies aimed at elucidating the electronic structures and catalytic cycles of these species. This chapter briefly reviews the most popular computational approaches employed in theoretical studies of bioorganometallic species and summarizes important information obtained from computational studies of (i) the enzymatic formation and cleavage of the Co–C bond of coenzyme B12; (ii) the catalytic cycle of methyl-coenzyme M reductase and its nickel-containing cofactor F430; (iii) the polynuclear active-site clusters of the bifunctional enzyme carbon monoxide dehydrogenase/acetyl-coenzyme A synthase; and (iv) the magnetic properties of the active-site cluster of Fe-only hydrogenases. KEYWORDS: bioorganometallic enzymes and cofactors carbon monoxide dehydrogenase/acetyl-coenzyme A synthase coenzyme B12 density functional theory electronic structure description Fe-only hydrogenase methyl-coenzyme M reductase

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1. INTRODUCTION Because of their complex geometric and electronic structures, the active sites and cofactors of bioorganometallic enzymes, which are characterized by the rare occurrence of metal–carbon bonds in Nature, represent major challenges for computational chemists. Nevertheless, owing to the remarkable progress in computer technology, the development of advanced computational methodologies, and the dedicated efforts of X-ray crystallographers that have afforded a wealth of high-resolution structures of the bioorganometallic enzymes and cofactors, the necessary foundation has been established for the successful completion of computational studies aimed at elucidating the electronic structures and catalytic cycles of these species. Density functional theory (DFT) has proven to be a particularly popular method for the quantum chemical treatment of these large metal-containing enzyme active sites and cofactors. The application of DFT in computational bioinorganic chemistry is the subject of an excellent review that has recently been published by Deeth [1]. As delineated in this chapter, computational studies have played a vital role in advancing our understanding of the electronic structures and catalytic cycles of the bioorganometallic enzyme active sites and cofactors. While the success in obtaining high-resolution X-ray structures has permitted detailed insight into the coordination environments of the metal centers in these species, in many cases delivering surprising information regarding the composition of polynuclear metal clusters and revealing the identities of unusual active-site ligands, these structures often raised more questions concerning the corresponding catalytic mechanisms than they answered. Therefore, computational studies – when properly evaluated on the basis of the results obtained in X-ray crystallographic, kinetic and/or spectroscopic investigations – will undoubtedly continue to play a key role in future research into the reaction cycles of the bioorganometallic systems. This chapter is organized as follows. In Section 2, the most popular computational approaches employed in theoretical studies of bioorganometallic species and methods for validating computational results on the basis of spectroscopic data are briefly reviewed. Section 3 summarizes relevant information obtained from computational studies of enzymes that are involved in the biosynthesis of coenzyme B12 or utilize this unusually complex cofactor for carrying out methyltransfer and radical-rearrangement reactions. The geometric and electronic properties of the nickel-containing cofactor F430 and the catalytic cycle of the enzyme that requires this species for its activity, methyl-coenzyme M reductase, are the subjects of Section 4. Section 5 provides an overview of important insights that have recently been gained from computational studies of the polynuclear active-site clusters of the bifunctional enzyme CO dehydrogenase/ acetyl-coenzyme A synthase. Lastly, in Section 6, a DFT-based quantitative

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analysis is presented of the magnetic properties of the active-site cluster of Feonly hydrogenases.

2. COMPUTATIONAL APPROACHES TO BIOORGANOMETALLIC CHEMISTRY 2.1. Overview The number of computational studies applied to bioorganometallic systems has grown so dramatically in the last decade that it is now almost commonplace to complement experimental studies with computations. Undoubtedly, a major contributor to this development has been the remarkable progress in computer technology, which is nicely demonstrated by the fact that the speed of microprocessor chips has increased by more than a factor of ten from 1997 to 2007 [2]. However, it would be a gross oversimplification to consider technological progress as the only driving force for the explosive growth of computational studies on bioorganometallic systems in the last decade. Computational approaches have been utilized with increasing frequency because they have proven to be an extremely useful complement to experimental investigations. For example, computations have been successfully used to investigate the formation and cleavage of the Co–C bond in cobalamin (Cbl, Section 3), evaluate putative catalytic intermediates in methyl-coenzyme M reductase (MCR, Section 4), explore the electron distributions and exchange interactions among the active site metal ions in carbon monoxide dehydrogenase/acetyl-coenzyme A synthase (CODH/ACS, Section 5), and elucidate the magnetic properties of the [FeFe] hydrogenase active-site cluster (Section 6). The foundation of computational approaches to chemistry is the prediction of energies and molecular geometries. There is a diverse selection of computational methods available to compute the electronic structure of a system at a fixed geometry. Below, these methods are categorized into distinct classes and the typical applications and limitations for each class are discussed using methylcobalamin (MeCbl) as a representative example (Section 2.2). All of these methods employ fundamentally similar strategies to find the energyminimized (or optimized) geometry of a bioorganometallic species. In particular, they all rely on the Born-Oppenheimer approximation [3], according to which the total wavefunction of a molecule is separated into electronic and nuclear components – a reasonable assumption considering that the electrons move much faster than the nuclei due to the large difference in mass. A typical geometry-optimization strategy can be illustrated by considering the Co–C bond of MeCbl [4]. First, a reasonable initial guess is made for the Co–C bond length and the total energy of MeCbl is computed for this particular nuclear configuration. Next, the Co–C bond length is changed slightly Met. Ions Life Sci. 2009, 6, 417–460


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and the total energy associated with this new geometry is computed. The geometry-optimization algorithm will continue to vary the Co–C bond length until it reaches a minimum for the total energy of the molecule. At this point, the geometry-optimization algorithm has optimized the Co–C bond length within the limitations of the method chosen to compute the total energy. In cases where the positions of all atoms of the molecule are simultaneously optimized, more complex algorithms are required [5], but the general conceptual approach remains the same. In general, the accuracy of the energyminimized (i.e., optimized) structure of the molecule is limited by the quality of the computational method used to calculate the total energy for a given nuclear configuration. It is of utmost importance to validate the molecular structure obtained in the energy minimization process on the basis of experimental data. In cases where detailed structural information about the molecule of interest is available, e.g., from X-ray crystallography or NMR spectroscopy, the accuracy of the optimized structure can be assessed from a direct comparison between key experimental and computed metric parameters. However, for structurally ill-defined species, such as short-lived intermediates, and to validate the computed electronic structure of a given molecule, it is useful to compare computationally predicted and experimentally determined spectroscopic parameters. A wide variety of methods have been developed to calculate spectroscopic observables from the computed electronic structure. Several methods that have been utilized to validate the computed electronic structure description for various Cbl species are described in Section 2.3. Since satisfactory agreement was achieved between theory and experiment for these species, the computational results provide a sensible framework for answering structural and mechanistic questions of interest.

2.2. Popular Methods for Computing Geometric and Electronic Structures Any cursory examination of the literature reveals that an enormous number of different methods is available for predicting the geometric structures of bioorganometallic species or chemical compounds in general. We do not attempt here to provide an exhaustive listing of all available methods or to present the quantitative details for any single method, as these topics have been thoroughly discussed in several text books [6–10]. Instead, this section highlights four methods that can be, and in fact have been, used to compute the geometric structures of bioorganometallic species (Table 1). Molecular mechanics (MM) is typically the method of choice to predict the structure of a large bioorganometallic species, such as a protein. MM is particularly appealing for studies of proteins because the large number of Met. Ions Life Sci. 2009, 6, 417–460

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LIPTAK, VAN HEUVELEN, and BRUNOLD Summary of the computational methods discussed in Section 2.2.

Method

Typical target

Molecular mechanics (MM) Density functional theory (DFT) Multiconfigurational self-consistent field (MC-SCF) Quantum mechanics/ molecular mechanics (QM/MM)

Protein Active site model Active site model valence MOs Active site (QM) +protein (MM)

Co–C bond description Harmonic spring Bonding MO Linear combination of MOs Depends on choice of QM method

Limitations Parameterization No electrons Single reference configuration Extreme computational cost Development of realistic QM/MM partitioning and coupling

atoms in these systems makes other currently available methods prohibitively expensive in terms of computational cost. The application of MM to protein systems has been recently reviewed [11,12]. A particularly popular implementation for bioorganometallic species is the Amber 95 force field [13,14]. Returning to the MeCbl example, this force field models the Co–C bond as a harmonic oscillator in which the two nuclei are connected by a spring. The equilibrium bond length and force constant of this spring are determined by parameters derived from a fit to reliable experimental or theoretical data. For example, the Amber 95 description of the Co–C bond of MeCbl required the careful development of the bond-length and forceconstant parameters for Co31 corrinoids by Marques and coworkers [15]. This extensive parameterization is a major limitation of MM, especially for bioorganometallic species where changes in the metal ion oxidation state and/or ligand environment due to substrate binding, product release, etc., will significantly affect the M–C bond length(s). A second major limitation of MM is that this method does not explicitly treat electrons and, thus, cannot be used for investigating the electronic structures of bioorganometallic species [16]. Density functional theory is often the method of choice for the explicit treatment of the electronic structure of a small bioorganometallic species, such as an active site model. Because DFT accounts for electron correlation in an approximate way, it is a more popular single reference configuration method than Hartree-Fock (HF) theory, which ignores this term altogether [10], for studying transition metal complexes where electron correlation is an important determinant of electronic structure [17]. The application of DFT Met. Ions Life Sci. 2009, 6, 417–460


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to bioinorganic species has been recently reviewed [1,18,19]. Note that in a DFT approach, the composition of every MO is simultaneously determined by minimizing the total energy of the molecule with the Kohn-Sham (KS) self-consistent field (SCF) method [20,21]. This description of the electronic structure as a single set of MOs is a significant limitation of DFT, especially for transition metal complexes where extensive mixing can occur between energetically proximate states that arise from different electronic configurations (so-called configuration interaction, CI) [22]. Multiconfigurational (MC)-SCF is usually the preferred method for properly treating CI mixing in transition metal complexes. If different electronic configurations give rise to multiple states that have similar total energies, then the electronic structure of the molecule cannot be accurately described by a single electronic configuration; rather, a mixture of these configurations must be considered. The application of the MC-SCF approach to transition metal complexes has been recently reviewed [22–24]. A relatively straightforward implementation of this approach is provided by the complete active space SCF (CASSCF) method [25]. CASSCF introduces multiconfigurational character into the ground-state wavefunction by allowing for CI mixing among the complete set of states arising from all possible electron configurations associated with a user-defined set of socalled ‘‘active’’ MOs. Because the CASSCF approach is intensely demanding of computational resources, only a relatively small number of active MOs can be considered. Consequently, the application of the CASSCF approach to bioorganometallic species requires that the user be extremely careful in selecting the proper set of active MOs. Combined quantum mechanics/molecular mechanics (QM/MM) is a computational strategy that employs a DFT or MC-SCF method for the treatment of a small portion of the molecule and an MM method for the remaining atoms. In principle, this is an ideal strategy for bioorganometallic protein systems, because it allows the electronic structure of the active site to be treated explicitly with DFT or MC-SCF, while the remainder of the protein can be modeled with MM to reduce the overall computational cost [16,26,27]. For example, when applied to a MeCbl-dependent enzyme, it would be sensible to describe the relevant portion of the cofactor and certain key amino-acid residues in the enzyme active site by a QM method and include all other atoms in the MM region to suitably account for steric and electrostatic interactions within the entire protein. The most challenging aspects of any QM/MM calculation are the partitioning of the full system into a QM region and a MM region and the treatment of the coupling between them. A proper treatment of the coupling terms is essential to assure that the MM region realistically perturbs the geometric and electronic structures of the QM region. The ideal partitioning and coupling of the QM and MM regions for a particular bioorganometallic species can require Met. Ions Life Sci. 2009, 6, 417–460

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significant user insight, but QM/MM ultimately offers an excellent strategy for accurately modeling the active sites of large enzymatic systems.

2.3. Computation of Spectroscopic Observables As stated above, it is of paramount importance that the results obtained with any theoretical method be validated on the basis of experimental data [18]. This is particularly true for computational studies of bioorganometallic species, whose electronic structures are too complex to be described by any currently available method without making certain approximations. The spectroscopic validation of computed electronic-structure descriptions entails the calculation of spectroscopic observables that can be compared directly to experimental data. Once satisfactory agreement between computed and experimental data is achieved, it is reasonable to trust the underlying electronic structure. The remainder of this section outlines some recent examples of how spectroscopic data were used as the basis for validating computational results obtained for the Cbl cofactor (Table 2). Electron paramagnetic resonance (EPR) spectroscopy offers one of the most sensitive probes of the ground state electronic structures of paramagnetic bioorganometallic species [28]. Therefore, several researchers have used EPR spectroscopy to investigate the paramagnetic cob(II)alamin (Co21Cbl) and cob(II)inamide (Co21Cbi1; a derivative of Co21Cbl in which the nucleotide loop including the DMB base is absent and a water molecule is axially coordinated to the cobalt ion) forms of the Cbl cofactor [29–32], and accurate values for both the g and 59Co A tensors could be extracted from quantitative analyses of the corresponding EPR spectra. The g and 59Co A tensors are sensitive to both the relative energies of the Co 3d-based MOs and the composition of the

Table 2. Summary of the spectroscopically validated calculations on the cobalamin cofactor discussed in Section 2.3.

Spectroscopy

Electronic structure method

Spectroscopic calculation method

Form of cobalamin

References

EPR Raman rR Abs Abs Abs Abs

B3LYP B3LYP PBE B3LYP BP86 PBE CASSCF

CPSCF SQM Harmonic TD-DFT TD-DFT TD-DFT CASPT2

Co21Cbl Co31Cbl Co11Cbl Co31Cbl Co21Cbl Co11Cbl Co11Cbl

[37] [52–54] [42] [41] [37] [42] [60]

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Co 3dz2-based singly occupied MO (SOMO), thus providing an excellent basis for assessing the feasibility of computational results. Hence, we have used the experimental EPR parameters for validating our DFT (B3LYP) computed [33–36] electronic-structure descriptions for Co21Cbl and Co21Cbi1 [37] by calculating the g and 59Co A tensors for each species with the coupled-perturbed SCF (CPSCF) method [38,39]. For both species the g and 59Co A tensors calculated by this method were found to agree reasonably well with the experimental data and, more importantly, accurately reproduced the nearly uniform increase of the g and 59Co A values from Co21Cbl to Co21Cbi1. This satisfactory agreement between the computationally predicted and experimentally observed trends in the g and 59Co A values indicated to us that the differences between the DFT computed relative energies and compositions of the Co 3d-based MOs of Co21Cbl and Co21Cbi1 are meaningful. Raman spectroscopy, which is excellently suited for probing the vibrational modes of bioorganometallic species [40], has been extensively utilized to study several biologically relevant forms of the Cbl cofactor [37,41–51]. Importantly, the vibrational frequencies obtained by Raman spectroscopy can be directly compared to the normal mode frequencies extracted from a calculated potential energy surface. Kozlowski and coworkers have taken advantage of this relationship and developed a DFT (B3LYP) based scaled quantum mechanical (SQM) force field for several forms of cob(III)alamin (Co31Cbl) [52–54] that was validated on the basis of experimental data. Although relatively poor agreement was achieved in the corrin stretching mode region (1450–1600 cm1), this force field satisfactorily reproduced the Co–C stretching mode region (350–650 cm1) of the Raman spectra of MeCbl and several other alkyl Co31Cbl species and, therefore, provided a suitable basis for exploring the origin of the different Co–C bond strengths in these species. More recently, we have been able to reproduce the corrin stretching mode region (1450–1600 cm1) of the Co11Cbl Raman spectrum almost quantitatively, without the need of introducing empirical scaling factors, by using the PBE density functional [34,42,55,56]. In particular, excellent agreement between theory and experiment was achieved for the vibrational frequencies, 1 H-2D isotopic shifts, and off-resonance Raman intensities. This study also attempted to predict the resonance Raman (rR) intensities of the corrinbased stretching modes. In contrast to the off-resonance Raman intensities, the rR intensities exhibit a strong dependence on the excitation wavelength used because only those normal modes that couple to the electronic transition at this particular wavelength are resonance enhanced [40]. We found that the PBE method successfully identified the resonance-enhanced modes in the corrin stretching mode region across the visible/near-UV regions of the Co11Cbl absorption spectrum. Owing to this good agreement between theory and experiment, a more detailed analysis of the DFT (PBE) computed bonding description for Co11Cbl was warranted. Met. Ions Life Sci. 2009, 6, 417–460

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Electronic absorption (Abs) spectroscopy has been the most frequently used spectroscopic tool in studies of the Cbl cofactor [57], in part because it can be utilized to probe electronic excited states regardless of the cobalt ion spin and oxidation states [58]. Although it is not typically possible to resolve the individual electronic transitions contributing to the Abs trace of a Cbl species directly, this task can usually be accomplished by collecting complementary data using circular dichroism (CD) and magnetic CD (MCD) spectroscopic techniques [37,41,42]. Collectively, the information that can be gained from a combined analysis of Abs, CD, and MCD spectroscopic data provides an almost ideal experimental framework for validating computational results. Thus, to assess the feasibility of the DFT computed electronic structure descriptions for our Co31Cbl, Co21Cbl, and Co11Cbl models, we have used the time-dependent DFT (TD-DFT) method [39,59] to predict the corresponding Abs spectra, which could then be compared directly to the results obtained from our Abs, CD, and MCD spectral analysis. Collectively, these studies revealed that DFT is well suited for obtaining reasonable electronic structure descriptions for all biologically relevant forms of Cbl. Jensen also used the Abs spectrum of Co11Cbl to validate his computed electronic structure description for this species [60], which he obtained by employing the CASSCF method [25]. In this study, the complete active space second-order perturbation theory (CASPT2) method was employed to compute the electronic transition energies and Abs intensities for Co11Cbl. Although the CASPT2 method successfully reproduced the energies of several electronic transitions observed experimentally, it failed to predict the correct number of electronic transitions contributing to the visible region of the Abs spectrum [42]. This example illustrates that while MC-SCF methods are generally more accurate than DFT methods, it may not be possible to include all of the relevant frontier MOs into the CASSCF active space for large bioorganometallic species due to current technological limitations.

3. FORMATION AND CLEAVAGE OF THE CO–C BOND OF COBALAMIN IN ENZYME ACTIVE SITES 3.1. Overview Vitamin B12 deficiency, which may be due to a number of factors, is a serious medical condition that causes megaloblastic anemia, hyperhomocystinuria, and methylmalonic aciduria [61,62]. In humans, these clinical symptoms can be triggered by defects in three distinct enzymes; namely, an ATP:corrinoid adenosyltransferase (ATR) [63,64], methylmalonyl-CoA mutase (MMCM) [65–67], and methionine synthase (MetH) [68,69]. ATR catalyzes the final step in the conversion of vitamin B12 to its physiologically active derivative Met. Ions Life Sci. 2009, 6, 417–460


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adenosylcobalamin (AdoCbl or coenzyme B12) by transferring the adenosyl (Ado) group from a molecule of adenosine-5 0 -triphosphate (ATP) to Co11Cbl. The AdoCbl product generated in this process is utilized by MMCM to catalyze the radical rearrangement of methylmalonyl-CoA (MMCoA) to succinyl-CoA (SCoA), which is a key step in the catabolism of branched-chain amino acids, odd-chain fatty acids, and cholesterol for entry into the Krebs cycle. Alternatively, MetH utilizes the MeCbl derivative of vitamin B12 for synthesizing methionine (Met) by transferring a methyl cation (CH+ 3 ) from methyltetrahydrofolate to homocysteine (Hcy). This reaction is essential for maintaining the proper distribution of cellular folate derivatives and preventing the accumulation of Hcy. Due, in part, to the fact that the reactions catalyzed by these three enzymes provide rare examples of organometallic transformations occurring in Nature, they have been the subjects of numerous computational investigations aimed at elucidating key steps in the corresponding catalytic cycles. The physiologically active forms of the Co31Cbl cofactor, AdoCbl and MeCbl, contain a low-spin Co31 ion that is equatorially ligated by four nitrogens from the corrin macrocycle and axially coordinated by a nitrogen from 5,6-dimethylbenzimidazole (DMB) on the a-face and a carbon atom from the Ado moiety or the CH3 group on the b-face (Figure 1) [57]. The formation of the axial Co–C bonds in AdoCbl and MeCbl involves the nucleophilic attack of a transient Co11Cbl species on the Ado moiety of ATP and the CH+ 3 group of a methylated substrate, respectively. Enzymes utilizing AdoCbl induce Co–C bond homolysis of the cofactor to produce Co21Cbl and an organic radical centered on the 5 0 -carbon of the Ado moiety (Adod) that is capable of abstracting a hydrogen atom from substrate as the first step in a protein-mediated substrate rearrangement reaction. Alternatively, the MeCbl-dependent enzymes catalyze methyl transfer reactions in which the cofactor’s Co–C bond is cleaved heterolytically, leaving behind a protein-bound Co11Cbl species. The remainder of this section is devoted to a discussion of how computational studies have enhanced our understanding of Co–C bond formation and cleavage by enzymatic systems.

3.2. Co–C Bond Formation During Adenosylcobalamin Biosynthesis and Methylcobalamin Activation by Methionine Synthase Since aquacobalamin is thought to be the circulating form of Cbl in the body, axial Co–C bonds of AdoCbl and MeCbl must be formed in vivo in reactions that are catalyzed by ATR and MetH, respectively. In each case a transient Co11Cbl species is generated in the enzyme active site that is a sufficiently strong nucleophile to abstract the Ado moiety of ATP or a methyl group Met. Ions Life Sci. 2009, 6, 417–460

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Figure 1. Chemical structure and numbering scheme for the physiologically active forms of Co31Cbl, AdoCbl and MeCbl. In aqueous solution, the coordination number of the Co ion is directly related to the oxidation state [106]. Upon reduction of Co31Cbl to Co21Cbl, the Co–X bond is cleaved. Further reduction of Co21Cbl to Co11Cbl also eliminates the axial Co–N bond. Note that in cobinamides the nucleotide loop and DMB base are absent and a water molecule is axially coordinated to the Co ion in the +3 and +2 oxidation states (corresponding to Co31Cbi1 and Co21Cbi1, respectively).

from adenosylmethionine (AdoMet). While these Co–C bond formation processes are relatively straightforward because Co11Cbl is one of the most potent nucleophiles known [71,72], the generation of an enzyme-bound Co11Cbl species via the reduction of Co21Cbl by a physiologically available Met. Ions Life Sci. 2009, 6, 417–460


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reductant poses a significant thermodynamic challenge. Even removal of the DMB moiety and its replacement by a solvent-derived water molecule (a weaker s-donor ligand) to produce base-off Co21Cbl is expected to raise the reduction potential by a mere 120 mV, to –490 mV versus SHE in aqueous solution (corresponding to the value reported for cob(II)inamide (Co21Cbi1) [73]). This value is still significantly more negative than the reduction potential of nicotinamide adenine dinucleotide phosphate (NADPH, –320 mV versus SHE [74]), the ultimate source of the reducing equivalents under physiological conditions. Therefore, ATR and MetH must have devised a different strategy for raising the Co21Cbl reduction potential to a sufficient degree that allows these enzymes to generate a Co11Cbl intermediate and, thus, to catalyze Co–C bond formation. Because the redox-active Co 3dz2-based molecular orbital of Co21Cbl is oriented along the axial coordination sites, which are singly occupied in Co21Cbl and vacant in Co11Cbl (Figure 1), it is to be expected that enzymatic manipulation of the Co21/11 reduction potential is accomplished by a destabilization of the Co21Cbl state rather than a stabilization of the Co11Cbl state. Indeed, Abs, MCD, and EPR spectroscopic data obtained for free and ATR(+ATP)-bound Co21Cbl exhibit significant differences, most notably the appearance of a prominent feature in the near-IR region of the MCD spectrum and widely spread resonances in the low-field region of the EPR spectrum when the cofactor binds to the enzyme active site. To interpret these changes in terms of geometric and electronic structural perturbations of Co21Cbl, we have used our spectroscopically-validated DFT computations as the basis for developing a spectro/structural correlation. Specifically, we have carried out a series of DFT and TD-DFT calculations to predict how distortions of the axial ligand–Co21 bonding interaction affect the Abs, MCD, and EPR spectra of Co21Cbl [37,75]. This correlation has enabled us to interpret the observed spectral changes accompanying the binding of Co21Cbl to ATR complexed with ATP in terms of the formation of an essentially four-coordinate Co21Cbl species that lacks any significant axial bonding interactions [75,76]. In this ‘‘activated’’ Co21Cbl species, the Co21/11 reduction midpoint potential is raised by an estimated 250 mV, which is sufficient to ensure that Co21-Co11 reduction can be accomplished in vivo. Using the same spectro/structural correlation for interpreting the Abs, MCD, and EPR spectroscopic data obtained for Co21Cbl bound to MetH locked into the cofactor activation conformation, we found that the Co–N bond on the a-face of the cofactor is broken and a water molecule is axially coordinated to the Co21 center on the b-face [77]. Furthermore, the axial Co–O(H2) bond of the MetH-bound Co21Cbl species is elongated relative to that of Co21Cbi1 (a model of base-off Co21Cbl, see Figure 1), presumably due to a hydrogen-bonding interaction with the phenolic group of the Y1139 Met. Ions Life Sci. 2009, 6, 417–460

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Figure 2. Enzyme-induced destabilization of the Co21Cbl state should raise the Co21Cbl reduction potential into a physiologically-accessible range.

residue that is located above the b-face [78]. Collectively, these perturbations to the axial coordination environment of Co21Cbl in MetH should be sufficient to raise the redox potential above that of Co21Cbi1 and thus into a physiologically accessible range (Figure 2).

3.3. Co–C Bond Homolysis in AdenosylcobalaminDependent Enzymes Of the different Co–C bond formation and cleavage processes discussed in this section, the enzyme-catalyzed Co–C bond homolysis of AdoCbl has by far received the most attention from the computational chemistry community. Since the last comprehensive review in 2001 [79], several research groups have engaged in theoretical studies that were aimed at elucidating the molecular mechanism of Co–C bond activation by AdoCbl-dependent enzymes [70,80–95]. The degree of Co–C bond activation accomplished by these enzymes is astonishing; in the case of MMCM, the rate of homolytic Co–C bond cleavage is elevated by as much as 1012-fold over that of the free cofactor [96]. This trillion-fold acceleration of Co–C bond homolysis by MMCM corresponds to a B17 kcal/mol reduction in the Co–C bond Met. Ions Life Sci. 2009, 6, 417–460


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dissociation enthalpy of AdoCbl in the enzyme active site, which could be achieved by a destabilization of the AdoCbl ‘‘ground state’’, a stabilization of the transition state, or both. Because Co–C bond homolysis is an endergonic process, the energy of the transition state could also be lowered indirectly, via a stabilization of the post-homolysis products Co21Cbl and the Adod radical, according to Hammond’s postulate. Therefore, enzymatic activation of the Co–C bond for homolysis can potentially be effected by any combination of AdoCbl destabilization, transition-state stabilization, and post-homolysis product stabilization. To explore how MMCM and a related enzyme, glutamate mutase (GM), activate the Co–C bond of AdoCbl for homolysis, we have carried out detailed spectroscopic studies of these enzymes and interpreted the corresponding data within the framework of spectro/structural correlations developed for AdoCbl and Co21Cbl [81–83]. While DFT computations predict that alterations in the axial bonding interactions or corrin fold angle should lead to fairly large shifts of the dominant electronic transition of AdoCbl, no such shifts were actually observed in our Abs, CD, and MCD spectra upon AdoCbl binding to MMCM or GM, even in the presence of substrate (analogues). Alternatively, rather significant differences in the MCD spectra of protein-bound versus free Co21Cbl were noted, especially in the region that is dominated by Co 3d-corrin p* charge transfer transitions. Analysis of these changes within the framework of DFT computations led us to propose that MMCM and GM induce a fairly uniform stabilization of the Co 3d-based MOs of Co21Cbl and, because these orbitals are filled, a stabilization of the enzyme-bound cofactor as a whole. As such, the results obtained in these combined spectroscopic and computational studies strongly suggest that stabilization of the post-homolysis product Co21Cbl, rather than destabilization of the AdoCbl ‘‘ground state’’, is a significant source of enzymatic Co–C bond activation. DFT (B3LYP) computations have also been used by Do¨lker and coworkers to investigate the effect of the MMCM active site on the Co–C bond homolysis rate of AdoCbl [85]. The computational model used in this study was derived from the crystal structure of MMCM [97] and included the full Ado moiety, the MMCoA substrate, and three active-site residues. MMCoA and the protein residues were frozen in their crystal structure orientations, while the AdoCbl geometry was optimized at three points along the Co–C bond dissociation coordinate. These geometry optimizations revealed that the post-homolysis products could be stabilized by a distortion of the Ado moiety that creates a favorable dipole-dipole interaction between the Co21 ion and two oxygen atoms of the Adod radical. This proposal is consistent with our hypothesis that product stabilization is a primary reason for the 1012-fold enhancement of the homolytic Co–C bond cleavage rate of AdoCbl by MMCM. Met. Ions Life Sci. 2009, 6, 417–460

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Most recently, two laboratories have utilized the QM/MM methodology to investigate the effects of the GM and MMCM active sites on the AdoCbl Co–C bond homolysis rate. Jensen and Ryde have used QM/MM computations to scan the potential energy surface of the Co–C bond of GM-bound AdoCbl from the ground state to the biradical product state [87]. These calculations confirmed that the Co21Cbl/Adod biradical product state is stabilized by an electrostatic interaction between the Co21 ion and two oxygen atoms from the Adod radical. They revealed further that an even larger contribution to the activation of the Co–C bond might involve the differential stabilization of the Ado moiety in the AdoCbl ‘‘ground state’’ and product states. Subsequent QM/MM calculations performed by Paneth and coworkers indicated that essentially the same strategy for Co–C bond activation of AdoCbl is employed by MMCM [92]. This study also successfully located the transition state along the Co–C bond dissociation coordinate. While the corrin ring conformation is minimally perturbed in the transition state, the Ado moiety is significantly distorted from its ground state structure (Figure 3). This result is consistent with the emerging view that the differential stabilization of the Ado moiety and stabilization of the Co21Cbl post-homolysis product are the primary contributions to the remarkable acceleration of Co–C bond homolysis by AdoCbl-dependent enzymes.

3.4. Co–C Bond Heterolysis in MethylcobalaminDependent Enzymes Compared to the Co–C bond homolysis step that is common to the catalytic cycles of all AdoCbl-dependent enzymes, the mechanism of enzyme-catalyzed Co–C bond heterolysis of MeCbl has received considerably less attention from the computational chemistry community [78,86,89,98–101]. While heterolytic Co–C bond cleavage in MeCbl formally produces a CH+ 3 cation (along with Co11Cbl), enzymes couple this process with substrate methylation; hence, the CH+ 3 cation does not actually represent an isolable intermediate [102]. This kinetic coupling complicates a theoretical analysis of the reaction profile of Co–C bond heterolysis by MeCbl-dependent methyltransferases, because both reactants, MeCbl and substrate, must evidently be considered when modeling this reaction. The intimate role that substrate plays in the methyl transfer reaction also suggests that there may not be a consensus mechanism employed by MeCbl-dependent methyltransferases. One of the best characterized members of this class of enzymes is MetH, which catalyzes a methyl group transfer from MeCbl to Hcy to produce Co11Cbl and Met [68,69]. MetH has been estimated to increase the rate of methyl transfer by 106-fold [103,104], with a 102-fold rate enhancement Met. Ions Life Sci. 2009, 6, 417–460


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Figure 3. The AdoCbl ground state (gray) and transition state (blue) of MMCM as computed by QM/MM. Reproduced from [92] with permission form the American Chemical Society, copyright (2006).

stemming from MeCbl binding and the remaining 104-fold acceleration arising from Hcy binding [105]. These results indicate that the mechanism employed by MetH to achieve the million-fold rate enhancement for Co–C bond heterolysis involves activation of both the cofactor and the substrate. Activation of the Met. Ions Life Sci. 2009, 6, 417–460

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Co–C bond of MeCbl could potentially entail an enzyme-induced elongation of the axial ligand–Co bond on the a-face (i.e., trans to the Co–CH3 bond, see Figure 1) of the cofactor, given that the preferred coordination numbers of the Co ion in MeCbl and Co11Cbl are six and four, respectively [106]. Computational support for this proposal was provided by a DFT (B3LYP) study performed by Do¨lker et al. in which the energy of Co–C bond heterolysis was found to decrease as a function of the trans Co–N bond length [86]. Moreover, evidence for the existence of a long axial ligand–Co bond opposite to the Co–CH3 bond has been obtained for at least one member of the family of methyltransferases. Specifically, in collaboration with other groups, we have performed a detailed characterization of the corrinoid/iron-sulfur protein (CFeSP), a MeCbl-dependent enzyme that binds the cofactor in the ‘‘base-off’’ form. A quantitative analysis of the spectroscopic data obtained for CFeSP within the framework of DFT and TD-DFT computations revealed that the axial Co–OH2 bond of CFeSPbound MeCbl is elongated by B0.2 A˚ relative to that of MeCbi1 [41,83,100]. This axial bond elongation should greatly facilitate the methyl group transfer from CFeSP-bound MeCbl to the ‘‘proximal’’ Ni center of the A-cluster of acetyl-coenzyme A synthase (see Section 5). Two additional strategies that could potentially be employed by MetH to enhance the rate of methyl transfer have been explored computationally by Jensen and Ryde [101]. This study utilized DFT (B3LYP) and a suitably truncated model system to compute the transition state for the methyl transfer from MeCbl to Hcy under several different reaction conditions (Figure 4). To justify the use of DFT in this study, the authors demonstrated that they achieved excellent agreement between the computed and experimental rates for the reaction in the absence of MetH by using the same computational methodology [103]. The first alternative strategy explored in this study involved Hcy deprotonation by enzyme-bound Zn [107,108]. Importantly, the DFT predicted activation energy for the methyl transfer reaction was found to be 22 kcal/mol larger for neutral Hcy than for its deprotonated (i.e., thiolate) form, suggesting that for the reaction to occur, substrate must first be deprotonated. The second strategy considered was the creation of a hydrophobic environment in the enzyme active site. The DFT computational results obtained by Jensen and Ryde [101] indicate that the lower dielectric constant expected for a hydrophobic active site can increase the rate of methyl transfer by up to 1014-fold. Therefore, the 106-fold rate enhancement by MetH could readily be achieved by deprotonation of Hcy and the creation of a hydrophobic active site. To this end, it is interesting to note that our spectroscopic studies of MetH locked into a catalytic conformation revealed that the active site is indeed inaccessible to solvent [75,77]. Met. Ions Life Sci. 2009, 6, 417–460


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Figure 4. The truncated model used in a DFT computational study of the methyl transfer reaction catalyzed by MetH. Reproduced from [101] with permission from the American Chemical Society, copyright (2003).

4. ORGANOMETALLIC CHEMISTRY AND CATALYTIC CYCLE OF METHYL-COENZYME M REDUCTASE 4.1. Overview Methyl-coenzyme M reductase (MCR) catalyzes the final step of methanogenesis in anaerobic archaea, a process that generates B109 tons of methane each year [109]. Crystallographic studies revealed that MCR is a 300 kDa hexamer of the form a2b2g2 that contains two equivalents of the F430 cofactor, which are separated by 50 A˚ [110]. This cofactor consists of a Ni center that is equatorially ligated by four nitrogens from the highly reduced hydrocorphin macrocycle and axially coordinated by O(Gln147) in the lower position and a variable ligand on the upper side (Figure 5) [111]. Access to the active site is controlled by a 30 A˚ channel that both excludes solvent and exerts steric control over the substrates, methyl-coenzyme M (Me-SCoM) and coenzyme B (HSCoB). In the first step of the catalytic cycle, Me-SCoM proceeds through the access channel and binds to the active site parallel to the F430 cofactor. Next, the thiol moiety of HSCoB is positioned approximately 8 A˚ above the Ni center by the amino acid residues along the channel [110]. The steric constraints imposed by the tunnel play a significant role in governing reactivity, as evidenced by the fact that enzymatic activity declines sharply when substrate analogues are used in which the length of the carbon chain differs from that of HSCoB [112]. Met. Ions Life Sci. 2009, 6, 417–460

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Figure 5. Top: Chemical structure of the cofactor F430 in the active Ni(I)MCRRed1 form of MCR, in which the lower axial position is occupied by O(Gln147) and the upper axial coordination site is vacant. Bottom: Chemical reaction catalyzed by MCR.

Numerous inactive forms of the enzyme, generated by the reaction of MCR with HSCoM and HSCoB, have been characterized and revealed that F430 can support +1, +2, and +3 oxidation states. Ni(II)MCRSilent is coordinated to the heterodisulfide CoM–S–S–CoB via the sulfonate oxygen of CoM, whereas Ni(II)MCROx1-Silent binds the thiol of CoM in the upper axial position of F430 [110,111]. Cryoirradiation of Ni(II)MCROx1-Silent yields NiMCROx1, which can be described as Ni(II) ion coupled to a thiyl radical in resonance with a Ni(III) thiolate [113]. Ni(I)MCRRed1, in which the upper axial position is vacant, serves as the active form of the enzyme. Met. Ions Life Sci. 2009, 6, 417–460


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Each form of MCR has unique spectroscopic signatures that provide a means of identifying the metal oxidation state and assessing the feasibility of computational models. The Ni(II)-bound forms of MCR exhibit a dominant Abs feature at 430 nm (F430 derives its name from this characteristic electronic transition) that undergoes a blue-shift to 378 nm upon reduction of the metal ion to the Ni(I)MCRRed1 state. MCD spectroscopic studies revealed that the Ni(I) form of the protein displays multiple low-energy transitions that are absent in the Ni(II)-bound states [113]. The vibrations of the tetrapyrrole ring, which can be probed by rR spectroscopy, also provide sensitive oxidation state markers, with key features downshifting upon nickel ion reduction [114]. Computational studies aimed at exploring the origin of these trends are discussed in Sections 4.2 and 4.3.

4.2. Properties of the Free F430 Cofactor Early computational studies of MCR focused on the isolated F430 cofactor, with the initial goals of identifying the structural and electronic characteristics that allow F430 to access multiple Ni oxidation states and evaluating the degree of ruffling in the cofactor. Nonplanar distortions are known to occur in a wide variety of tetrapyrrole-type species, both in natural systems such as heme and in synthetic complexes, and such distortions can significantly influence the corresponding electronic properties [115,116]. In 1991, Farber et al. crystallized a pentamethylester derivative of the native cofactor [117]. When F430 is isolated from MCR, it converts to its diepimer form, eventually reaching an equilibrium of 88% conversion to the catalytically-inactive and highly ruffled 12,13-diepimer (F430Diepi) [118]. Unlike the diepimer, native F430 is relatively planar [119] and has Ni–N bonds that are longer than those of its ruffled counterpart (1.99–2.14 A˚ in F430 as compared to 1.96–1.99 A˚ in F430Diepi) [118]; these longer bonds in turn stabilize the Ni(I) oxidation state. Similarly, the longer Ni–N bonds in the native cofactor are more conducive to high spin Ni(II) than the smaller low spin Ni(II) ion [118]. DFT geometry optimizations of the native cofactor and F430Diepi accurately reproduced this trend in Ni–N bond lengths [110]. The planar geometry necessary for the catalytic function is preserved by the steric influence of amino acid residues within the MCR active site, a finding supported by both DFT and molecular mechanics studies [118–121]. Computational data also support the somewhat counterintuitive claim that F430 can stabilize the Ni(III) oxidation state, even though it favors high-spin Ni(II) over its smaller low-spin counterpart. DFT studies by Wondimagegn and Ghosh [119] of [Ni(III)(F430)(ClO4)(Am)]1 and [Ni(III)(F430)(CH3)(Am)]1 (Am ¼ O-bound acetamide) revealed that the upper axial ligand determines the nature of the singly occupied Ni 3d orbital. The methyl moiety, which is a Met. Ions Life Sci. 2009, 6, 417–460

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strong s-donor, results in a preferential (3dx2y2)1 electron configuration, consistent with EPR studies of MCROx1 [122], whereas the perchlorate ligand favors a (3dz2)1 electron configuration. Further, a geometry optimization of the [Ni(III)(F430)(CH3)(Am)]1 model resulted in Ni–N bond lengths similar to those found in F430, as expected by considering that in each case the Ni 3dx2y2based orbital is singly occupied. Collectively, these results indicate that the long Ni–N distances favored by the planar cofactor are crucial for supporting oxidation states ranging from Ni11 to Ni31 [123]. Because of the large size of F430, truncated models have typically been used in computational studies of this species. To assess the minimal model size needed for accurately reproducing the structural and electronic properties of the complete F430 cofactor, Pelmenschikov et al. [124] performed DFT computations on two differently truncated models. This study revealed that a relatively small model with a simplified 12-carbon macrocycle is sufficient to preserve the key electronic properties of F430. However, a larger model in which the pyrrole and lactam rings were preserved and the propionate and acetate groups replaced with hydrogen atoms was needed to properly describe the steric influences imposed by neighboring amino acid residues and subtle conformational changes in the cofactor during catalysis [124,125]. Consistent with these findings, our DFT and TD-DFT computations for models of F430 that contained the pyrrole rings accurately reproduced the experimental Abs spectra and EPR g and A tensors, even when the lactam ring was replaced with hydrogen atoms [126]. Collectively, the results obtained in these studies demonstrate that the pyrrole rings must be preserved in computational models of F430 in order to accurately reproduce experimental observations, whereas truncations of peripheral side chains and even the lactam ring are relatively unproblematic. Additionally, these studies established DFT as a reliable method for computing structures, energies, and electronic properties of the F430 cofactor. The results obtained in DFT studies of the free F430 cofactor also permitted us to develop an MO-based explanation for an intriguing spectroscopic property of this species; namely, that the dominant Abs feature in the visible spectral region undergoes a rather dramatic blue-shift from 430 to 380 nm upon reduction of nickel ion from the +2 to the +1 oxidation state [126]. On the basis of our spectroscopically-validated bonding descriptions obtained from DFT computations on Ni(II)F430 to Ni(I)F430 models, this feature can be assigned to a hydrocorphin-centered p-p* transition. In the oxidized Ni(II)F430 species, the Ni-based 3d orbitals are considerably lower in energy than the p-based frontier orbitals of the macrocycle and thus do not participate in electronic transitions contributing to the visible region of the Abs spectrum. However, upon one-electron reduction of the Ni(II) ion, the occupied Ni 3d orbitals are raised in energy, shifting between, and strongly mixing with, the hydrocorphin p and p*-based MOs in Ni(I)F430. Met. Ions Life Sci. 2009, 6, 417–460


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These ground-state changes have a dramatic effect on the excited-state structure, causing a blue-shift of the dominant p-p* transition and the appearance of numerous Ni 3d-hydrocorphin p* charge-transfer features in the visible/near-IR region that give rise to a series of prominent features in the corresponding MCD spectrum [126]. Hence, Abs and MCD spectroscopic studies are well suited for assessing the Ni oxidation state in each of the plentiful known states of MCR-bound F430, especially when combined with EPR experiments.

4.3. Ni-Alkyl Species of Methyl-Coenzyme M Reductase: Computational Evaluation of Viable Reaction Mechanisms The rapid kinetics associated with the reaction catalyzed by MCR has thus far precluded the observation of any intermediates. Therefore, models of viable reaction intermediates have been prepared using substrate analogues and characterized by spectroscopic methods, and computations have been carried out to evaluate possible reaction mechanisms. These studies led to the proposal of two fundamentally different mechanisms for the catalytic cycle of MCR. Mechanism 1 involves a Ni(III)–Me intermediate, whereas mechanism 2 instead invokes the intermediacy of a methyl radical and a Ni(II)–thiol species [127].

4.3.1. Ni–Methyl Pathway Analogous to known cobalamin chemistry (Section 3.4), mechanism 1 invokes an organometallic intermediate. Here, Ni(I) performs a nucleophilic attack on the methyl moiety of Me-SCoM, resulting in a putative Ni(III)–CH3 intermediate [128,129]. Two variants of this mechanism have been suggested in the literature. In the scheme put forward by Ragsdale and coworkers on the basis of steady-state kinetic data, the first step in the reaction involves the generation of a SdCoB radical species [129]. The second variation, favored by Thauer and coworkers (Figure 6, top), instead invokes the intermediacy of an S–CoB anion [128]. According to this latter mechanism, residues in the substrate access channel, namely Tyr333 and Tyr367, serve as proton donors to the reactants at key steps along the reaction pathway. Both of these mechanisms are consistent with structural data. Recent studies of MCR using substrate analogues revealed that the formation of alkyl–Ni(III)F430 species is chemically viable. Specifically, the reaction of Ni(I)MCRRed1 with bromopropanesulfonate (BPS) was found to yield a Ni(III)–propyl sulfonate species, termed MCRPS, that has EPR properties Met. Ions Life Sci. 2009, 6, 417–460

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Figure 6. Proposed reaction mechanisms for MCR. Adapted and combined from [125,128].

similar to MCROx1 [130]. Similarly, the reaction of Ni(I)MCRRed1 with CH3X (X ¼ Br, I) was shown to generate a Ni(III)–CH3 species known as MCRMe [131,132]. EPR and electron-nuclear double resonance (ENDOR) studies established that, like [Ni(III)(F430)(CH3)(Am)]1 (discussed in Section 4.2), MCRMe has a (3dx2y2)1 ground state [131]. Both alkyl-Ni(III)MCR species mimic the reactivity of native reaction intermediates of MCR; Ragsdale and coworkers demonstrated that MCRMe can be reacted with HSCoB to produce methane [131], and MCRPS can be treated with a thiolate to regenerate Ni(I)MCRRed1 [133]. However, theoretical studies have questioned the viability of mechanism 1. Crabtree and co-workers used DFT computations to quantify the energy Met. Ions Life Sci. 2009, 6, 417–460


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barrier associated with the initial reaction step; namely, the breaking of the C–S bond in Me-SCoM to form an alkyl–Ni(III) species. For the two forms of the putative organometallic intermediate considered, CH3–Ni(II)F430 and CH3–Ni(III)F4301, DFT yielded Ni–C binding energies of 24.9 and 18.0 kcal/ mol, respectively. In comparison, the C–S bond strength in Me-SCoM was calculated to be 70 kcal/mol [124]. On the basis of these results, it would be expected that the methyl transfer from Me-SCoM to Ni(I)MCRRed1 is highly endothermic, by at least B45 kcal/mol. This finding led to the proposal of mechanism 2 described next [124].

4.3.2. Methyl Radical Pathway The obvious shortcomings associated with mechanism 1 led to the suggestion that the nucleophilic attack of Ni(I)MCRRed1 on Me-SCoM causes the C–S bond to cleave homolytically (as opposed to heterolytically, as implicated in mechanism 1) to generate a methyl radical and a CoM–S–Ni(II)F430 species (Figure 6, bottom). Methane is then produced through hydrogen atom abstraction from HSCoB, a process that generates a SdCoB radical species that subsequently reacts with CoM–S–Ni(II)F430 to form the mixed disulfide CoM–S–S–CoB and restore Ni(I)MCRRed1 [118,124,125,134]. Mechanism 2 was evaluated computationally by Siegbahn and coworkers [124,125]. Importantly, the computed Ni–S bond strength of 39 kcal/mol for the putative CoM–S–Ni(II)F430 species that is formed in this process is much closer to the 70 kcal/mol needed to cleave the S–C bond in Me-SCoM (vide supra). In further support of mechanism 2, the DFT computed energy barrier associated with this mechanism is only 20 kcal/mol and the overall reaction was found to be exothermic [124,125]. A somewhat troubling aspect of mechanism 2 is that it relies on the direct transfer of the methyl radical from Me-SCoM to HSCoB over a relatively large distance. Although the F430 cofactor and the active-site tunnel may provide the necessary steric restrictions for the controlled transfer of a reactive radical to occur, both substrates are held in place through a series of hydrogen bonds, such that their thiol groups are separated by B6.5 A˚ [133]. Another potential problem with mechanism 2 is that the driving force associated with the disulfide bond formation between two distant thiol moieties may not be sufficient to drive the Ni–S bond cleavage in the putative CoM–S–Ni(II)F430 species [134]. Lastly, it should be noted that experimental studies of MCR using the substrate analogue ethyl-SCoM revealed that an inversion of configuration at the reactive carbon occurs on reduction; thus, if a methyl radical is indeed formed in the MCR catalytic cycle, then it must be quenched by HSCoB on a timescale that is sufficiently short to prevent it from rotating [127,135]. Met. Ions Life Sci. 2009, 6, 417–460

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4.3.3. Protonation of Ni(I)MCRRed1 Recently Duin and McKee [136] proposed a third type of mechanism for MCR based on a DFT computational study. This mechanism relies on the protonation of Ni(I)MCRRed1, at either the Ni(I) site or the tetrapyrrole ring, followed by oxidative addition of Me-SCoM. The disulfide bond between MeSCoM and HSCoB is formed through a two-center three-electron interaction between the two thiol moieties, leaving the liberated methyl group on the protonated NiF430 cofactor. Protonation of the methyl group then leads to release of methane and regeneration of Ni(I)MCRRed1. Some novel aspects of this mechanism, such as the displacement of Ni from the tetrapyrrole ring by 1.5 A˚ and the coordination of Ni to both the thiol and the methyl of MeSCoM, bear further experimental and computational investigation [136].

5. GEOMETRIC AND ELECTRONIC STRUCTURES OF THE CARBON MONOXIDE DEHYDROGENASE/ ACETYL-COENZYME A SYNTHASE ACTIVE SITE CLUSTERS 5.1. Overview The bifunctional enzyme CO dehydrogenase/acetyl-coenzyme A synthase (CODH/ACS) is crucial for the autotrophic growth of some archaea and eubacteria. The CODH/ACS from the anaerobic mesophile Moorella thermoacetica (formerly called Clostridium thermoaceticum) [137], which is probably the best studied member of this class of enzymes, is a 310 kDa tetramer of configuration a2b2 [138,139]. Each ab dimer contains four polynuclear metal clusters that are labeled A- through D-clusters [140–142]. The B-, C-, and D-clusters reside in the b-subunit and are involved in the reversible reduction of CO2 to CO (equation 1) for subsequent generation of acetyl-CoA at the A-cluster. The B- and D-clusters each consist of relatively typical [Fe4S4] cubanes and act as a ‘‘redox wire’’, shuttling the electrons necessary for the reduction of carbon dioxide between the C-cluster and external reducing agents. The C-cluster itself contains an [Fe3S4] core that is connected to a Ni–S–Fe moiety (Figure 7, bottom). Remarkably, the CO generated at the C-cluster via CO2 reduction passes to the A-cluster through a 138 A˚ intramolecular tunnel within CODH/ACS [143]. 2e þ 2Hþ þ CO2 Ð CO þ H2 O

ð1Þ

The C-cluster can be stabilized in multiple oxidation states, including Cox, Cred1, Cred2, and Cint. The electron distribution among the multiple redox-active sites of the C-cluster in these states remains incompletely understood. Met. Ions Life Sci. 2009, 6, 417–460


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Nevertheless, spectroscopic studies revealed that Cox is an EPR-silent (presumably S ¼ 0) species, as is Cint. In contrast, both Cred1 and Cred2 are S ¼ 1/2 systems that can be readily distinguished on the basis of their characteristic g tensors. Redox titrations demonstrated that the one-electron reduction of Cox generates Cred1, which can undergo sequential one-electron reductions to yield first Cint and then Cred2 [144–146]. The A-cluster, found in the a subunit, catalyzes the synthesis of acetylCoA according to the following reaction: CH3 -CFeSP þ CO þ CoA Ð acetyl-CoA þ CFeSP

ð2Þ

where CH3-CFeSP and CFeSP are the methylated and de-methylated forms, respectively, of the corrinoid/iron-sulfur protein (see Section 3.4). Two

Figure 7. (bottom).

Chemical structures of the CODH/ACS A-cluster (top) and C-cluster

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relatively recent X-ray crystallographic studies [147,148] revealed that the Acluster consists of an [Fe4S4] cubane that is linked through a cysteine bridge to a tetrahedral, proximal Ni site (Nip), which in turn is connected to a square-planar, distal Ni site (Nid) via two cysteine bridges (Figure 7, top). While initially the presence of two nickel centers at the A-cluster came as a surprise, it was soon recognized that these structures provided the necessary framework for understanding previously un-interpretable spectroscopic data. Additionally, they provided definitive clues as to the origin of the inherent heterogeneity of the A cluster, alternately showing Zn and Cu (in addition to Ni) at the proximal metal site, which is therefore referred to as the labile site. Subsequent studies afforded compelling evidence that Ni is present in the active form of ACS [145,147,148]. Collectively, spectroscopic studies on CODH/ACS revealed that the Acluster is stable in at least three states: the as-isolated oxidized state (Aox) in which both metal centers are low-spin Ni(II), a one-electron reduced state (Ared), and a CO-bound, one-electron reduced state (Ared–CO) [137]. Conversion of Aox to the Ared state leads to the development of an S ¼ 3/2 EPR signal that is characteristic of a reduced [Fe4S4]11 cubane, indicating that the reducing equivalent localizes on the FeS cluster. The Ared–CO state exhibits the well-characterized S ¼ 1/2 NiFeC EPR signal, so named for the hyperfine broadening associated with labeled 61Ni, 57Fe, and 13CO [149]. The relevance of this Ared–CO species to catalysis is debated (see Section 5.3).

5.2. Mechanism of Carbon Monoxide Oxidation/Carbon Dioxide Reduction by Carbon Monoxide Dehydrogenase The mechanism of CODH is most commonly discussed as the oxidation of CO, as many enzymes, such as the Mo/Cu-dependent CODH and the monofunctional Ni-dependent CODH (which lacks the a-subunit and thus the ability to synthesize acetyl-CoA formation), generate CO2. As carbon monoxide metabolism was recently reviewed [145], we provide only a brief overview here. Although the complex geometric and electronic structures of the C-cluster greatly complicate experimental studies of reaction intermediates, it is generally believed that CO and H2O bind to the Ni(II) and Fe(II), respectively, of the Ni– S–Fe moiety in the Cred1 state (note that this Fe center is referred to as ferrous component II (FCII, Figure 7, bottom) because of an early Mo¨ssbauer study [140]). The Fe–OH species then attacks the Ni21–CO moiety to form a Ni21– COOH intermediate, which rapidly decays by the release of CO2 and the elimination of a proton to yield Cred2. The transfer of two electrons through the ‘‘redox wire’’ regenerates Cred1 while reducing the B- and D-clusters, which in Met. Ions Life Sci. 2009, 6, 417–460


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turn are oxidized by ferredoxin to return the b subunit to the ‘‘ready’’ state [111,145,146]. Until recently, the structure and composition of the C-cluster were subjects of considerable debate, which precluded computational studies of the CODH reaction mechanism. However, the high-resolution X-ray crystal structures of the C-cluster that were recently determined [147,150] have afforded the necessary foundation for the successful completion of future computational studies aimed at elucidating the catalytic cycle of CODH at the molecular level.

5.3. Computational Insights into the Catalytic Cycle of Acetyl-Coenzyme A Synthase Studies of the catalytic cycle of ACS are complicated by the intricate, multimetallic nature of the active site A-cluster. Computational studies have been instrumental in identifying the metal in the proximal site of the catalytically active A-cluster and in discriminating between putative catalytic cycles. As mentioned in Section 5.1, early X-ray crystal structures of CODH/ACS showed Zn, Cu, and Ni in the proximal metal site of the A-cluster [145,147,148]. To determine which metal is catalytically active, we conducted a DFT study that considered two putative models of Ared–CO (note that even though this species may not actually be catalytically relevant, the intensity of the NiFeC EPR signal correlates directly with enzyme activity [151]), one containing Cu and the other Ni in the proximal site [149]. In each case, a CO moiety was bound to the proximal metal site and DFT was used to carry out a constrained geometry optimization in which the positions of certain C atoms of ligating amino acid residues were kept frozen to account for the constraints imposed by the protein backbone. As both models yielded structures consistent with experimental data for the A-cluster, the DFT/ ZORA CP-SCF method was used to compute the corresponding 57Fe, 61Ni, and 13CO nuclear hyperfine parameters. It was found that only the model containing Ni in the proximal site successfully reproduced all of the experimental Aiso, indicating that the NiFeC species, and thus the catalytically active A-cluster form possesses a proximal nickel ion [149]. The NiFeC species (i.e., Ared–CO state of the A-cluster) can therefore be described as 2+ core. possessing a [Fe4S4]21–Ni+ p CO–Nid It is widely accepted that the C-cluster generates CO that subsequently migrates to the A-cluster and that CFeSP delivers a methyl cation to the A-cluster. Additionally, because treatment of the A-cluster with 1,10-phenanthroline (phen) abolishes catalytic activity by removing the labile Nip center, it is generally assumed that Nip serves as the primary site of substrate CO binding [152]; however, this is the limit of consensus regarding the mechanism of ACS. There are two general classes of proposed mechanisms to date; namely, the paramagnetic and diamagnetic pathways. In the paramagnetic mechanism the Met. Ions Life Sci. 2009, 6, 417–460

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EPR-active Ared–CO state serves as an early intermediate that binds the methyl group from CH3-CFeSP to generate a Nip(CO)(CH3) species. Migratory insertion of CO into the Ni–CH3 bond then yields a Ni–acetyl intermediate, and attack of this species by CoAS– leads to the formation of the product acetyl-CoA cycle, [145,153]. In an alternate proposal that also employs a proximal Ni3+/1+ p methylation precedes carbonylation [154]. Some variants of the paramagnetic mechanism involve the cycling of the iron-sulfur cluster between +1 and +2 states [153]; however, experimental evidence has cast doubt on this proposal, as kinetic studies revealed that the [Fe4S4]21/11 redox conversion is orders of magnitude slower than the observed reaction rate of the A-cluster [155]. An alternative to the paramagnetic mechanism is a diamagnetic scheme, in which CH+ 3 binds first to the proximal Ni center and all intermediates are EPRredox cycling of the proximal nickel ion and thus silent, assuming a Ni2+/0+ p invoking an unprecedented zero-valent Ni center. This description arises from 2+ the apparent two-electron reduction of Aox, a [Fe4S4]21-Ni2+ p -Nid species, to generate the catalytically active form of the A-cluster [156]. Additionally, Riordan and coworkers recently demonstrated that a methylated cobalt species, MeCo(dmgBF2)2py (dmgBF2¼(difluoroboryl)dimethylglyoximato and py ¼ pyridine), can transfer a –CH3 moiety to a Ni0 model compound, a reactivity that mirrors the Ni2+/0+ diamagnetic proposal [157]. p As with the F430 cofactor, theoretical studies of the A-cluster must carefully consider the degree of active site truncation that can be used without compromising the accuracy of computational results. Webster et al. [158] employed a model of the A-cluster in which the [Fe4S4] cubane was modeled by a mononuclear high-spin Fe(II) tetrathiolate. Interestingly, DFT computations performed on this model seemed to lend support to the diamagnetic mechanism over the alternative Ni3+/1+ pathway [158]. However, two Ni2+/0+ p p independent studies by us and Field et al. demonstrated that the redox-active [Fe4S4]21/11 cluster must be included in its entirety in order to obtain accurate computational results [149,159]. In particular, inclusion of the complete iron-sulfur cluster in DFT computations caused an initial [Fe4S4]21–Ni0+ p – 2+ model to optimize to a [Fe4S4]11–Ni1+ species in which the ironNi2+ d p –Nid sulfur cluster and proximal Ni are antiferromagnetically coupled, producing an overall diamagnetic species in accordance with experimental data [149]. The fact that the computed electron distributions within the A-cluster vary considerably depending on the size of the active site model used clearly warrants additional computational studies of this highly elaborate metal cluster. Field and coworkers [159] performed a series of DFT studies to evaluate proposed reaction pathways; namely, they considered the possibility of the distal Ni center acting as a binding site (the so-called binuclear pathway [153]), the order in which CH+ 3 and CO bind to Ni, and the stability of the putative zerovalent nickel center. While their findings did not definitively discriminate between initial methylation versus carbonylation, they found the structures Met. Ions Life Sci. 2009, 6, 417–460


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associated with a mononuclear mechanism, in which both CO and CH+ 3 bind to Nip, to be energetically more favorable than a binuclear mechanism involving both the proximal and distal Ni centers [159]. Their findings suggest that Nid remains in the +2 oxidation state throughout the reaction, in good agreement with experimental data. Additionally, their results revealed that while the reduced [Fe4S4]11 cubane may aid in the stabilization of the zero-valent state of the proximal Ni center, in its oxidized state (which is invoked in the diamagnetic mechanism as required by experimental data) the [Fe4S4]21cluster oxidizes Ni0+ p to Ni1+ p , consistent with our findings described above [149,159].

6. MAGNETIC PROPERTIES OF THE ACTIVE SITE CLUSTER OF IRON-ONLY HYDROGENASES 6.1. Overview Central to hydrogen metabolism in aerobic and anaerobic microorganisms is a family of enzymes known as hydrogenases that catalyze the consumption and production of H2 via the following reversible reaction [160,161]: H2 Ð 2Hþ þ 2e

ð3Þ

There are three known classes of hydrogenases, which differ in their activesite compositions: the nickel-iron ([NiFe]), iron-only ([FeFe]), and iron-sulfur cluster-free hydrogenases [162]. The [NiFe] and [FeFe] hydrogenases are primarily used for dihydrogen oxidation and proton reduction, respectively, while the iron-sulfur cluster-free enzymes activate H2 for use in catabolic processes. The catalytic mechanisms of the [NiFe] and [FeFe] hydrogenases have been the subjects of numerous DFT computational studies. For a summary of the results obtained in these studies, we refer the reader to an excellent review by Siegbahn, Tye, and Hall that has very recently been published [162]. Instead, this section focuses on a topic not covered elsewhere in this chapter; namely, the use of computations for elucidating the magnetic properties of a polynuclear active-site cluster. Specifically, we summarize here our computational efforts to model the exchange interactions that operate within the active-site cluster of the [FeFe] hydrogenases.

6.2. Structure and Oxidation States of the Active Site Cluster of [FeFe] Hydrogenases While not as widespread as [NiFe] enzymes, the [FeFe] hydrogenases have attracted a great deal of interest due, largely, to their unusual 6-Fe active-site cluster known as the H-cluster. X-ray crystallographic studies revealed that Met. Ions Life Sci. 2009, 6, 417–460

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the H-cluster consists of a [Fe4S4]H cubane linked to a [2Fe]H subcluster (Figure 8) [161,163]. Each iron atom of the [2Fe]H component is terminally coordinated by one CN and one CO ligand, and the two Fe centers are bridged by a dithiolate ligand and an additional CO ligand (COb). The proximal Fe (Fep) is connected to the [Fe4S4]H cubane via a cysteine bridge, while the distal Fe (Fed) has a labile coordination site (L) that is the putative site of H-binding during catalysis [162]. During catalysis, the [2Fe]H component likely cycles between Fe(II)-Fe(II) and Fe(I)-Fe(I) states, making [FeFe] hydrogenases the only known biological system to employ the Fe(I) oxidation state [164–166]. The active form of the H-cluster is the paramagnetic (S ¼ 1/2) Hox state in which the [2Fe]H component possesses a mixed-valence {Fep-Fed}31 configuration. The Hox state readily binds CO, a potent inhibitor of [FeFe] hydrogenases, at the Fed center to generate the Hox-CO form, while treatment with dithionite yields the EPR-silent Hred state that features an {Fep-Fed}21 unit. The [Fe4S4]H cubane is formally in its diamagnetic +2 state in all well-characterized forms of the H-cluster; however, spectroscopic studies have shown that in the Hox and Hox-CO forms the four Fe sites of the cubane display 57Fe hyperfine coupling parameters (A values) that are diagnostic of extensive spin delocalization from the [2Fe]H site onto the cubane [167–170]. Nearly all theoretical investigations of the H-cluster performed to date have omitted the [Fe4S4]H cubane and instead modeled this moiety by adding a proton to the Cys residue bound to Fep [171–174]. While this approach may be justified by the fact that the [2Fe]H subunit is presumably the site of H2

Figure 8.

Structure of the active site H-cluster of [FeFe]-hydrogenases.

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uptake/formation, spectroscopic evidence suggests that a high degree of electronic communication exists between the two units, and it is thus likely that the [Fe4S4]H cubane influences both the geometric and electronic structures of the [2Fe]H component. To address these issues, we employed DFT to perform the first computational investigation of the H-cluster in its entirety and subsequently performed a detailed analysis of the magnetic properties of the Hox and Hox-CO states using DFT methods [175]. Importantly, our geometry optimizations of complete and truncated H-cluster models revealed that the positions of the SCys and COb ligands are strongly correlated, as the shorter Fep–SCys bonds in models lacking the [Fe4S4]H cubane caused the COb ligand to shift away from Fep by B0.10 A˚.Thus, it appears that an important structural role of the [Fe4S4]H cluster is to lengthen the Fep–SCys bond, which in turn modulates the relative position of the COb ligand. This finding raises some concerns as to the credibility of previous computational studies that omitted the [Fe4S4]H cubane from H-cluster models when investigating the catalytic mechanism of [FeFe] hydrogenases.

6.3. Calculation of the Magnetic Properties of the H-Cluster The primary difficulty in treating the complete H-cluster with DFT arises from the exchange interactions that operate between the two subclusters as well as within the [Fe4S4]H cubane. Even though the cubane as a whole is diamagnetic, it actually consists of two pairs of high-spin Fe21 (S ¼ 2) and Fe31 (S ¼ 5/2) centers [176]. Double-exchange coupling yields two ferromagnetically coupled (S¼9/2) mixed-valence pairs of Fe centers, which in turn couple antiferromagnetically to produce the S ¼ 0 ground state observed experimentally. The H-cluster can thus be modeled as a three-spin system with SI ¼ SII ¼ 9/2 and SH ¼ 1/2, where SI and SII represent the spins of the two mixed-valence Fe dimers that comprise the cubane, and SH represents the spin associated with the [2Fe]H subunit in the Hox and HoxCO states (Figure 9) [175]. Consequently, this model requires two exchange parameters, Jcube and JH, to describe the Heisenberg exchange coupling between the two Fe dimers within the cubane and between the [2Fe]H component and the adjacent Fe dimer, respectively. Using this spin-coupling model along with the broken-symmetry (BS) approach developed by Noodleman and coworkers [177,178], Jcube and JH can be computed from the relative energies of the high-spin and BS states, as indicated in Figure 9 (note that because in DFT wave functions are represented by single determinants, only the high-spin (S ¼ 19/2) state can be properly described by this method; however, the energies of the other pure spin states, and thus the Jcube and JH values, can be calculated from the energies of the BS states [175]). Met. Ions Life Sci. 2009, 6, 417–460

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Figure 9. Left: Schematic diagram of the ‘‘three-spin model’’ used to treat the H-cluster. Right: Spin state diagram showing how the relative energies of the high-spin and brokensymmetry (BS) states of the [Fe4S4]2+ H cluster are modulated by ferromagnetic (F) and antiferromagnetic (AF) interactions with the S ¼ 1/2 spin of the [2Fe]H component.

The computed Jcube values of B400 cm1 for both the Hox and Hox-CO states are consistent with previous computational [177,178] and experimental [179] studies of [Fe4S4]21 clusters. More importantly, the calculated JH parameter of +15 10 cm1 for the Hox state agrees remarkably well with the experimental value of B20 cm1 determined from Mo¨ssbauer studies [167,168]. Our computations also successfully reproduced the large increase in JH upon binding of CO to Fed, predicting a JH value of 15050 cm1 for the Hox-CO state that closely matches the experimental value of B100 cm1 [167,168]. Given this good agreement between the experimental and computed exchange-coupling parameters, it was reasonable to use the electronic-structure descriptions provided by DFT as the basis for exploring the origin of the large increase in JH upon CO-binding to the H-cluster [175]. This analysis revealed that in the Hox state, the unpaired spin density is almost entirely localized on the Fed center, which lacks an effective pathway for exchange coupling to the [Fe4S4]H cubane. Alternatively, the large amount of unpaired spin density on Fep in the Hox-CO state facilitates exchange interactions with the cubane via the bridging Cys residue. Thus, the extent of delocalization of unpaired spin density over the Fe dimer determines the magnitude of the exchange coupling to the [Fe4S4]H cluster, and the JH value therefore serves as a useful indicator of the spin distribution within the [2Fe]H component.

7. CONCLUDING REMARKS AND FUTURE DIRECTIONS Collectively, the remarkable progress in computer technology, the development of advanced computational methodologies, and the dedicated efforts of enzymologists, spectroscopists, and X-ray crystallographers have established Met. Ions Life Sci. 2009, 6, 417–460


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the necessary foundation for the successful completion of computational studies aimed at elucidating the electronic structures and catalytic cycles of the bioorganometallic enzymes and cofactors. Although the majority of the computational studies completed to date relied on DFT, a stable and relatively reliable method for the quantum chemical treatment of large metalcontaining species, it can be anticipated that more sophisticated theoretical tools will be employed with increasing frequency in future studies of bioorganometallic species. Among these, the combined QM/MM and MC-SCF approaches appear particularly well suited for evaluating viable catalytic cycles by utilizing complete protein models and for elucidating the magnetic properties of exchange-coupled polynuclear active-site clusters, respectively. Additionally, it is to be expected that theoretical chemists will continue to make improvements in the computational prediction of spectroscopic observables, thus establishing a more rigorous framework for evaluating calculated bonding descriptions, structures of hypothetical reaction intermediates, and viable catalytic mechanisms on the basis of experimental data. Computational studies have played a vital role in elucidating key steps in the catalytic cycles of enzymes involved in the biosynthesis of the B12 cofactors as well as of AdoCbl-dependent enzymes (Section 3). In the case of the Co–C bond-forming ATP:corrinoid adenosyltransferases, combined spectroscopic/computational studies revealed that these enzymes effect the thermodynamically difficult Co21-Co11 reduction through the formation of an essentially square-planar Co21corrinoid intermediate, so as to stabilize the redox-active, Co 3dz2-based molecular orbital that is oriented along the axial coordination sites of the Co21 ion. A consensus has also been largely reached regarding the mechanism of Co–C bond activation by AdoCbldependent enzymes; namely, that the differential stabilization of the Ado moiety and stabilization of the Co21Cbl post-homolysis product are the primary contributors to the remarkable acceleration of Co–C bond homolysis by these enzymes. In contrast, because the MeCbl-dependent enzymes have received much less attention from the computational chemistry community, relatively little is known to date about the mechanism of enzymecatalyzed Co–C bond heterolysis. Likewise, despite numerous kinetic, spectroscopic, and computational studies, the reaction mechanism of MCR is an enduring subject of intense debate (Section 4). Substrate-analogue studies have revealed that the formation of alkyl–Ni(III)F430 species is chemically viable and that such species can react with HSCoB to generate methane and a disulfide, even as theoretical studies have cast doubt on the possibility of forming such an organometallic intermediate in the reaction with the native substrate Me-SCoM. Computational methods have also been used with great success in studies of the ACS A-cluster (Section 5) and the [FeFe] hydrogenase H-cluster (Section 6). In both cases, DFT has been utilized to assign the oxidation states of Met. Ions Life Sci. 2009, 6, 417–460

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individual metal ions within the active-site clusters and to evaluate key steps in the corresponding catalytic cycles. However, nearly all theoretical investigations of the A-cluster and H-cluster performed to date have omitted the [Fe4S4] cubane. While this approach may seem reasonable because the substrates bind to other metal centers in these clusters, spectroscopic and computational evidence suggests that the [Fe4S4] cubanes significantly influence both the geometric and electronic structures of the remaining portions of the active sites. Consequently, future computational studies aimed at elucidating the catalytic mechanisms of ACS and [FeFe] hydrogenases should employ Acluster and H-cluster models that include the complete [Fe4S4] cubane. Lastly, because the structure and composition of the CODH C-cluster have recently been determined by X-ray crystallography, the necessary foundation has also been laid for future computational studies to evaluate the proposed catalytic cycle of CODH at the molecular level.

ACKNOWLEDGMENTS T.C.B. thanks his current and former graduate students and postdoctoral fellows for their hard work and valuable discussions on this project, his superb collaborators for generous protein supply, Professor Frank Neese for providing us with a free copy of his ORCA computational software package and for his advice on electronic structure calculations, and acknowledges the National Science Foundation (CAREER award MCB-0238530) for financial support. K.M.V.H. was supported by the National Science Foundation Graduate Research Fellowship Program.

ABBREVIATIONS Abs ACS Ado AdoCbl AdoMet Am ATP ATR B3LYP BP86 BPS

electronic absorption acetyl-coenzyme A synthase adenosyl adenosylcobalamin S-adenosylmethionine acetamide adenosine-5 0 -triphosphate adenosyltransferase Becke’s three-parameter hybrid functional for exchange coupled with the Lee-Yang-Parr correlation functional nonlocal gradient corrections of Becke for exchange and Perdew for correlation bromopropanesulfonate

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BS CAS CASPT2 Cbi Cbl CD CFeSP CI CODH CP DFT DMB dmgBF2 ENDOR EPR FCII Fed Fep Gln GM Hcy HF HSCoB HSCoM IR KS MC MCD MCR Me MeCbl Me-SCoM Met MetH MM MMCM MMCoA MO NADPH Nid Nip PBE

453

broken symmetry complete active space complete active space second-order perturbation theory cobinamide cobalamin circular dichroism corrinoid iron-sulfur protein configuration interaction carbon monoxide dehydrogenase coupled-perturbed density functional theory 5,6-dimethylbenzimidazole (difluoroboryl)dimethylglyoximato electron-nuclear double resonance electron paramagnetic resonance ferrous component II of the C-cluster in CODH/ACS distal metal site in the H-cluster of [FeFe]-hydrogenase proximal metal site in the H-cluster of [FeFe]-hydrogenase glutamine glutamate mutase homocysteine Hartree-Fock coenzyme B coenzyme M infrared Kohn-Sham multiconfigurational magnetic circular dichroism methyl-coenzyme M reductase methyl methylcobalamin methyl-coenzyme M methionine methionine synthase molecular mechanics methylmalonyl-CoA mutase methylmalonyl-CoA molecular orbital nicotinamide adenine dinucleotide phosphate distal metal site of the A-cluster in CODH/ACS proximal metal site of the A-cluster in CODH/ACS Perdew-Burke-Ernzerhof generalized gradient approximation Met. Ions Life Sci. 2009, 6, 417–460

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phen py QM rR SCF SCoA SHE SOMO SQM TD ZORA

1,10-phenanthroline pyridine quantum mechanics resonance Raman self-consistent field succinyl-CoA standard hydrogen electrode singly-occupied molecular orbital scaled quantum mechanical time-dependent zeroth order regular approximation

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Subject Index

A Absorption spectroscopy cobalamins, 426, 429, 431 F430 model, 438, 439 methyl-coenzyme M reductase, 437 UV, see UV absorption spectroscopy Acetaldehyde, 100 phosphono-, 83 Acetate (or acetic acid), 59, 72, 76, 162 Acetamide, 437, 438, 440 Acetobacterium dehalogenans, 77, 80 Acetogenesis, 35 Acetonitrile, 117, 119 Acetyl-coenzyme A, 31, 59, 73, 77, 136, 139, 451 biosynthesis, see Biosynthesis decarbonylase/synthase, 136, 140 Acetyl-coenzyme A synthase(s), 31, 434, 443 active site, see Active sites catalytic cycle, 445–447, 452 density functional theory calculations, see Density functional theory calculations EPR studies, see EPR mechanism, 146, 445, 446 models, 446 nickel-carbon bonds, see Nickel-carbon bonds

Acetyl-coenzyme A synthase/carbon monoxide dehydrogenase, 133–146, 419, 420 A-cluster, 135–141, 144, 146, 442–444 active site, see Active sites azide in, see Azide B-cluster, 135, 136, 146, 442 catalytic properties, 137, 138 C-cluster, 134–138, 141–146, 442 cyanide in, 135, 142–144 D-cluster, 135, 136, 442 diamagnetic mechanism, 137, 138, 140 electron density, see Electron density electron transfer in, see Electron transfer ENDOR studies, see ENDOR spectroscopy EPR studies, see EPR from Moorella thermoacetica, 133–146, 442 hydrogen bonds, see Hydrogen bonds hydrolysis, 141 infrared studies, see Infrared spectroscopy inhibitors, 142, 143 methyl transfer, see Methyl transfer Mo¨ssbauer spectroscopy studies, see Mo¨ssbauer spectroscopy nickel-carbon bonds, see Nickel-carbon bonds paramagnetic mechanism, 140, 141 ping-pong mechanism, 140, 141 redox properties, 137, 138 structure, 135, 443, 444


462 Acidity constants (see also Equilibrium constants), 385, 399 B12 derivatives, 20 H2, 236 ACS/CODH, see Acetyl-coenzyme A synthase/carbon monoxide dehydrogenase Active site (of) acetyl-coenzyme A synthase, 445, 446 ACS/CODH, 135, 136, 442–447 blue copper proteins, 338 bromoperoxidase, 369 carbonic anhydrase, 385 catechol dioxygenase, 380 catechol oxidase, 303 cobalamins, 426–435 copper amine oxidase, 334 copper-zinc superoxide dismutase, 331 cytochrome c oxidase, 318, 324 deoxyhemocyanin, 300 dopamine b-monooxygenase, 308, 311, 315–317 F430, 125–127 [FeFe]-hydrogenases, 153, 161, 179–208, 222, 231, 447–449 galactose oxidase, 337 glutamate mutase, 88, 432 heme, 261, 266 heme-copper oxidases, 317–319, 326 hemocyanin, 303, 308 hydrogenases, 152 manganese catalase, 375 methyl-coenzyme M reductase, 115–129, 435, 437 methylmalonyl-coenzyme A mutase, 431, 432 MoCu CODH, 333 multi-copper oxidase, 339 [NiFe]-hydrogenases, 153, 155–165, 168–172, 200, 222, 231 nitrite reductases, 341, 342 nitrous oxide reductase, 343 non-heme iron enzymes, 379 peptidylglycine a-hydroxylating monooxygenase, 311, 315–317 protocatechuate 3,4-dioxygenases, 382 superoxide reductase, 381 tyrosinase, 303 xanthine oxidoreductases, 398, 400, 403, 406, 409–413 zinc enzymes, 384, 386

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX Acyl-coenzyme A synthase, 57–60, 73, 76 decarbonylase, 57, 73, 74 Adenosine 5 0 -deoxy-, 84, 85, 91 5 0 -diphosphate, see 5 0 -ADP 5 0 -triphosphate, see 5 0 -ATP Adenosylcobalamin(s) (see also Coenzyme B12 and Vitamin B12), 18, 38, 54, 83, 84, 86, 87, 90, 91, 104–106, 427, 428, 430–432 3 0 ,4 0 -anhydro-, 97 as cofactor, 34, 40, 96, 102 base-off, 93 biosynthesis, see Biosynthesis DBM-off/His-on, 103 DBM-on, 96, 101 5 0 -deoxy-5 0 -, 3 -dependent rearrangements, 84–106 His-on, 102 reactivity, 24 structure, 4, 11, 12, 14, 55, 88, 98 S-Adenosylmethionine, 34, 61, 62, 65, 67, 68, 75, 79, 80, 81, 83, 84, 93, 428 radical, see Radicals Adenosyltransferase, 34, 92, 93 chaperone, see Chaperones 5 0 -ADP, 59 Affinity constants, see Stability constants Agaricus, 304 bisporus, 307 Alcaligenes sp., 341 Alcohols, see individual names Alcohol dehydrogenase zinc in, 383 Aldehyde(s) (see also individual names), 334, 3335, 337 acet-, see Acetaldehyde glycol, 84 hydration, 385 oxidase, 368 propion-, 84 Aldehyde oxidoreductase, 398, 403, 411 crystal structure, see Crystal structures Aldolase fructose-1,6-diphosphate, 383 Algae (see also individual names), 181, 184, 373 brown, 369 green, 183 marine, 368, 372

SUBJECT INDEX [Algae (see also individual names)] oxygen evolving complex, see Oxygen evolving complex red, 246, 369 Alkaline phosphatases, 386, 388 magnesium, 388 zinc, 288 Alkylation, 26 Co(I), 25 Co(III)-corrins, 31 cob(I)amides, 34 cobalamins, 25, 28, 30, 77 Allochromatium vinosum, 158 Allopurinol as inhibitor, 404 Alloxanthine xanthine oxidoreductase complex, 404–408 Amide bond cleavage, 386 nickel coordination, 136, 139 proton exchange, 269 Amine(s) (see also individual names) aromatic, 339 oxidation, 334, 335 tris(pyrid-2-ylmethyl)-, see Tris(pyrid-2ylmethyl)-amine Amino acids (see also individual names) radicals, see Radicals Amino acid sequences [FeFe]-hydrogenases, 186, 203 [Fe]-hydrogenases, 223 heme oxygenase, 278, 279 olfactory receptors, 346 Aminomutases, 38, 95–103 lysine, see Lysine 2,3-aminomutase ornithine, see Ornithine aminomutase Aminopeptidase (see also Peptidases and individual names), 386 leucine, see Leucine aminopeptidase Ammonia, 100, 101, 370–373 aspartate lyase, 202 Amyotrophic lateral sclerosis, 330 Anemia hemolytic, 245 megaloplastic, 426 Antibacterial agents (see also individual names), 340 Antibiotics (see also individual names), 83, 346–349 biosynthesis, see Biosynthesis

463 Anticancer agents (see also individual names), 347, 387 Antiglaucoma agents, 387 Antiinflammatory drugs, see Drugs effect of carbon monoxide, 245 Antiosteoporosis agents, 387 Antioxidants (see also individual names), 246, 247 Apoptosis inhibition, 245 Aquacobalamin, 54, 66, 427 Co(III), 3, 19–21, 23, 29, 79, 81, 93, 100 electrochemistry, 20, 26 structure, 4, 9, 10, 13 Arabidopsis sp., 345 thaliana, 246 Archaea (see also individual names), 54, 56, 60, 84, 165, 181, 221, 442 anaerobic, 58, 134, 435 methanogenic, 73, 83, 116, 221, 234, 386 methanotrophic, 116 Arene Cu(I) complex, 298, 306, 307 Arrhenius plot, 23 Arthrobacter globiformis, 334, 336 Arthropods (see also individual names) hemocyanin, 299, 301, 302, 304–306 Ascophyllum nodosum, 369, 372 Ascorbate, 308, 339 as reductant, 312 Ascorbate oxidase, 339 rate constants, see Rate constants structure, 340 zucchini squash, 339 Aspartate transcarbamoylase, 383 Aspartic acid methyl-, 86, 87 Association constants (see also Stability constants) carbon monoxide binding, 257, 258 dioxygen binding, 257, 258 heme oxygenases, 257, 258 Atmosphere carbon monoxide in, 243, 332 50-ATP, 59, 77, 79–81, 92, 93, 162, 371, 373, 429 biosynthesis, see Biosynthesis corrinoid adenosyltransferase, see Transferases hydrolysis, 75, 82, 373

Met. Ions Life Sci. 2009, 6, 461–496

464

SUBJECT INDEX

[50-ATP] production, 327 -using adenosyltransferase, 34 ATPase, 100 Azide (in), 255, 366, 373, 375–377, 387 ACS/CODH, 135, 142 chloroperoxidase, 369 heme oxygenase, see Heme oxygenases as inhibitor, 269, 270, 274, 376, 377 manganese catalase, 377 manganese superoxide dismutase, 376 superoxide reductase, 381, 382 Azotobacter chroococcum, 370, 371 nitrogenase, 370 vinelandii, 370 Azurin, 338

B Bacteria(l) (see also individual names), 56, 161, 165, 181, 231 acetogenic, 59, 73, 74 anaerobic, 34, 78, 134, 181 cyano-, see Cyanobacteria eu-, see Eubacteria heme oxygenase, see Heme oxygenases hyperthermophilic, 184 Knallgas, 171 oxygen evolving complex, 381 pathogenic, 246, 248, 251 photosynthetic, 282, 283 purple non-sulfur, 202 purple, 381 soil, 380 sulfate-reducing, 152, 165, 166, 171, 184 sulfidogenic, 78 thermophilic, 134, 184 Benzimidazole 5,6-dimethyl-, see 5,6-Dimethylbenzimidazole 5-hydroxy-, 70, 74, 80, 81 5-methoxy-, 54 Benzoate chlorinated, 78 Bilirubin, 246 Biliverdin, 245–247, 251, 253, 256, 268 Fe(II), 247 Fe(III), 247 formation, 276–282, 284

Met. Ions Life Sci. 2009, 6, 461–496

Binding constants, see Association constants, Equilibrium constants, and Stability constants Bioorganometallic species computational studies, 419–452 Biosynthesis 13 C-labeled heme, 260 acetyl-coenzyme A, 35, 443 adenosylcobalamin, 427–430 antibiotics, 378 ATP, 317 biotin, 204 carbon monoxide, 241–285 coenzyme B12, 3, 34, 419 [Fe]-hydrogenases, 227 FeMo cofactor, 202, 204 fosfomycin, 82 H-cluster of [FeFe]-hydrogenases, 199–207 iron guanylylpyridinol cofactor, 225 lipoic acid, 204, 205 melanin, 299 methionine, 35 Biotechnology, 366 Biotin biosynthesis, see Biosynthesis synthase, 203 Bleomycin, 346–349 Co(III), 347 Cu(I), 346–349 Cu(I)-carbon monoxide, 348 Cu(II), 347–349 cytotoxicity, 349 Fe(II), 346–348 hydroperoxo-Fe(III), 347 mechanism of action, 347 metal-free, 347–349 NMR studies, see NMR structure, 347, 348 Blue copper proteins (see also individual names), 337, 338 active site, see Active site electron density, see Electron density electron transfer in, see Electron transfer EPR studies, see EPR redox potentials, see Redox potentials Bonds amide, 386 C–C, 33 C–O, 403 C–S, 441 CH3–S, 125, 127

SUBJECT INDEX

465

[Bonds] Co–C, see Cobalt-carbon bonds Cu–C, see Copper-carbon bonds Cys–His, 304 dissociation energies, 31, 32, 38 hydrogen, see Hydrogen bonds Mo–C, 402, 404 Mo=O, 397–399, 411 Mo=S, 397, 400, 401, 408 Ni–C, see Nickel-carbon bonds Ni(II)–S, 124 O–O, 252, 255, 256, 267, 268, 379 thioether, 304 Born-Oppenheimer approximation, 420 Bos taurus (see also Bovine) xanthine oxidoreductase, 398 Bovine copper-zinc superoxide dismutase, 330, 331 cytochrome c oxidase, 318, 320, 326, 329 heart heme-copper oxidase, 318, 321, 322,327 xanthine oxidoreductase, see Xanthine oxidoreductase Brain, 245 Bromide methyl-, see Methyl bromide oxidation, 372 Bromoperoxidases active site, see Active site inhibition, 372 vanadium, 369, 372 3-Bromopropane sulfonate, 118, 120–122, 439 [3-13C]-, 121 as inhibitor, 120 Buffer Tris, see Tris(hydroxymethyl)methylamine Busycon canaliculatum, 302, 308 carica, 301 Butyrate 4-bromo-, 122 4-hydroxy-, 122

C Cadmium(II) in HybD protease, 164 Calorimetry photoacoustic, 320, 321

Camberus loevis, 305 robustus, 305 Cancer (see also Carcinomas and individual names) ovarian, 347 risk assessment, 28 testicular, 347 Cancer irroratus, 305 Carbamates, 385 thio-, see Thiocarbamates Carbamoyl phosphate, 201 Carbamoyl phosphate synthase, 161, 162 Carbohydrates (see also individual names), 373 fermentation, 182 metabolism, see Metabolism Carbon 11 C-labeling, 350 12 C-labeling, 233, 331 13 C ENDOR, see ENDOR 13 C NMR, see NMR 13 C-labeling, 96, 101, 123, 139, 146, 158, 162, 225, 259, 260, 300, 331, 397, 400, 444, 445 14 C-labeling, 139 -carbon bond, 23, 86 -molybdenum bond, see Bonds nickel bond, see Nickel-carbon bonds -oxygen bond, see Bonds skeleton rearrangements, 85–95 -sulfur bond, see Bonds Carbon dioxide (in), 58, 60, 72, 73, 76, 77, 221, 298, 332, 385, 387 13 CO2, 162 ACS/CODH, 134, 143–145 fixation, 35 hydration, 384 reduction, 56, 57, 373, 442, 444, 445 Carbonic anhydrase (from) active site, see Active sites catalysis of hydrolytic processes, 385 classes, 386 crystal structure, see Crystal structures human, 387 hydrogen bonds, see Hydrogen bonds inhibitors, 387 iodide complex, 387 mammalian, 385 mechanism of action, 385 Methanosarcina thermophila, 385

Met. Ions Life Sci. 2009, 6, 461–496

466 [Carbonic anhydrase (from)] plant, 385 zinc in, 383–387 Carbon monoxide (in), 57, 73, 75, 77, 137, 142–144, 146, 297, 366, 367, 373, 442, 445, 447 12 CO, 233 13 CO, 139, 225, 444, 445 as inhibitor, see Inhibition as probe in copper amine oxidase, 336 atmospheric levels, 243, 332 biosynthesis, see Biosynthesis Cu(I) binding, see Copper(I) cytoprotective effects, 243–245 dehydrogenase, see Carbon monoxide dehydrogenases dopamine b-monooxygenase, 311–316 environmental sources, 243 Fe(II)-CO, see Iron(II) [FeFe]-hydrogenases, 179–208, 448, 450 [Fe]-hydrogenases, 219–237 heme oxygenases, see Heme oxygenases heme-copper oxidases, 319–325 hemes, see Hemes metabolism, see Metabolism [NiFe]-hydrogenases, 158, 160–164, 172 nitrite reductase, 341, 342 oxidation, 59, 134, 202, 298, 444, 445 peptidylglycine a-hydroxylating monooxygenase, 311–316 photodissociation, 302 properties, 243 release from heme, see Heme role in biology and medicine, 244 stretching frequencies, 305, 315, 349 toxicity, see Toxicity Carbon monoxide dehydrogenases, 57, 59, 73, 75, 80, 202, 398, 444, 445 catalytic mechanism, 146, 445 electron density, see Electron density ferredoxin, 445 from Carboxythermus hydrogenoformans, 142 from Rhodospirillum rubrum, 136, 141 inhibitors, 142, 143 MoCu, see Molybdenum-copper carbon monoxide dehydrogenase nickel insertion, 164 X-ray diffraction spectroscopy studies, see X-ray diffraction spectroscopy

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX Carbon monoxyhemocyanin, 304–308 mutation, 307 Carboxydothermus hydrogenoformans, 59, 73–75, 136, 142 Carboxylase pyruvate, see Pyruvate Carboxylate groups, 375 bridging, 376 Carboxypeptidases (see also Peptidases), 386 A, 383 Carcinoma (see also Cancer) head, 347 neck, 347 skin, 347 Catalases, 374 manganese, see Manganese catalase Catecholase activity, 299, 304 Catechol dioxygenases, 377, 378, 380 extradiol-cleaving, 380 Fe(II)-active site, 380 intradiol-cleaving, 380 Mn(II)-active site, 380 Catechol oxidase (from), 298–300, 306 active site, see Active sites crystal structure, see X-ray crystal structure Ipomoea, 303 reaction cycles, 303, 304 Cell damage, 244 death, 327 proliferation, 245 smooth muscle, 245 Cephalosporin, 378 biosynthesis, see Biosynthesis Ceruloplasmin, 339, 340 CD, see Circular dichroism Chaperones (for) adenosyltransferase, 93 bacterial copper CusF, 344, 345, 350 diol dehydrase, 100 flavodoxin, 81 methyltransferase-activating protein, 82 Charge transfer, 439 metal-to-ligand, 306, 307 Chemotherapy, 347, 404 Chlamydomonas reinhardtii, 183 Chloride Ni(II) complex, 119 oxidation, 372

SUBJECT INDEX Chlorinated compounds dechlorination, 77, 78 detoxification, see Detoxification Chloroform, 119, 122 Chloroperoxidase azide in, see Azide vanadium, 368, 369 Chromatium vinosum, see Allochromatium vinosum Chromatography high performance liquid, see High performance liquid chromatography hydrophobic interactions, 228 Chromophores, 284 Ciliates (see also individual names), 184 Circular dichroism (studies of) cobalamins, 426, 431 far-UV, 346 Citrate, 235 homo-, see Homocitrate titanium, 72 Clostridium barkeri, 90 pasteurianum, 152, 181–192 stricklandii, 102, 103 thermoaceticum, see Moorella thermoacetica Clusters (in) [7Fe-9S-homocitrate], 370 [8Fe-7S], 370 [Fe2S2], see [Fe2S2] cluster [Fe3S4], see [Fe3S4] cluster [Fe4S4], see [Fe4S4] cluster [Mn(III)Mn(III)], 375, 376 2Fe sub-, see [FeFe]-hydrogenases ACS/CODH, see Acetyl-coenzyme A synthase/carbon monoxide dehydrogenases Cu4S, 343 cubane (see also [Fe4S4] cluster), 136, 139–141, 442–444, 448, 449 H-, see [FeFe]-hydrogenases iron-sulfur, 34, 35 Mn4Ca, 374 nitrogenases, 370 oxygen-bridged binuclear non-heme iron, 378 P-, 370

467 Cobalamins (see also Coenzyme B12, Corrinoids, Vitamin B12, and individual names) absorption spectroscopy, see Absorption spectroscopy active site, see Active sites adenosyl-, see Adenosylcobalamin adenylylpentyl-, 100, 105 alkylation, see Alkylation aqua-, see Aquacobalamin base-off, 5, 12, 14, 15, 20, 21, 26, 74–76, 78, 79, 81 base-off/His off, 62, 63 base-off/His on, 39, 62, 63, 67, 74, 77, 79, 90, 93 base-on, 5, 7, 12, 14, 19, 20, 22, 26, 29, 39, 74, 76, 106 chloro-, 4 circular dichroism studies, see Circular dichroism Co(I), 8, 19, 20, 22, 28, 29, 32, 37, 61, 65, 66, 77, 78, 87, 92, 93, 424–430, 432, 434 Co(II), 5, 9, 10, 15, 19, 20, 29, 31–34, 38–40, 61, 62, 65–67, 76–79, 81, 82, 84, 85, 87, 88, 90–93, 96, 97, 106, 424–432, 451 Co(III), 9, 10, 13, 19, 22, 26, 29, 34, 54, 66, 79, 424–426, 428 cyano-, see Cyanocobalamin density functional theory calculations, see Density functional theory calculations -dependent enzymes, 53–107 electron density, see Electron density electron transfer in, see Electron transfer epi-, 7, 8, 13, 26 EPR studies, see EPR EPR studies, see EPR His-off, 65, 66 His-on, 65, 66, 86 hydrogen bonds, see Hydrogen bonds hydroxo-, see Hydroxocobalamin magnetic circular dichroism studies, see Magnetic circular dichroism methyl transfer, see Methyl transfer methyl-, see Methylcobalamin nitroxyl-, 4 propyl-, 66, 67 Raman spectroscopy studies, see Raman spectroscopy

Met. Ions Life Sci. 2009, 6, 461–496

468 [Cobalamins (see also Coenzyme B12, Corrinoids, Vitamin B12, and individual names)] spectroscopically validated calculations, 424 structure, 4, 9, 55, 428 trichlorovinyl-, 77, 78 vinyl-, 12, 35 Cobalt (oxidation state undefined) 59 Co, 424, 425 -carbon bonds, see Cobalt-carbon bonds Cobalt(I) (in) alkylation, see Alkylation B12 derivatives, 18–20, 26 cobalamins, see Cobalamins corrinoids, see Corrinoids corrins, see Corrins redox couples, see Redox potentials Cobalt(II) (in), 72 B12 derivatives, 18–20 cobalamin, see Cobalamins cobinamides, see Cobinamides cobyrinate, see Cobyrinate corrinoids, see Corrinoids corrins, see Corrins methylcobalamin, see Methylcobalamin redox couples, see Redox potentials reduction, 451 Cobalt(III) (in), 1–41, 138 bleomycin, see Bleomycin cobalamins, see Cobalamins cobamides, see Cobamides cobinamides, see Cobinamides cobyrinate, see Cobyrinate corrinoid iron-sulfur protein, see Corrinoid iron-sulfur protein corrinoids, see Corrinoids corrins, see Corrins methylcobalamin, see Methylcobalamin redox couples, see Redox potentials Cobalt-carbon bonds, 145 activation, 84 cleavage, 24, 30–34, 84, 87, 91, 106, 420, 430–435 formation, 24–31, 426–435, 451 heterolytic cleavage, 31, 36, 432–435, 451 heterolytic formation, 30, 36 homolytic cleavage, 31, 32, 38–40, 96, 97, 100, 430–432, 451 homolytic formation, 30 reactivity, 3

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX Cobamides, 4 adenosyl-, 4, 33 alkylation, see Alkylation base-off, 82 base-off/His-on, 70 base-on, 54, 82 Co(II), 74–76, 82 Co(III), 34, 38, 74 5,6-dimethylbenzimidazole-, see Cobalamins 5-hydroxybenzimidazolyl-, 70, 74, 80, 81 imidazolyl-, 7, 12, 13, 26, 33, 36, 37 5 0 -methoxybenzimidazole, 54 methyl transfer, see Methyl transfer methyl-, 74, 82 p-cresolyl-, 18, 55, 56 structure, 8, 9, 15 Cobesters (see also Corrinoids), 5 Co(II), 5, 15, 30 structure, 6 Cobinamides (see also Corrinoids and Vitamin B12) adenosyl-, 14, 34 aqua-, 82 base-off, 75 Co(I), 21, 23, 32, 76 Co(II), 21, 32, 33, 75, 82, 424, 428, 429 Co(III), 21, 428 density functional theory calculations, see Density functional theory calculations diaqua, 21 EPR studies, see EPR methyl-, see Methylcobinamides structure, 6, 428 Cobyrinate, 5, 13 Co(II), 26 Co(III), 13, 14 structure, 6 CODH, see Carbon monoxide dehydrogenases Coenzyme A, 57, 73, 91 acetyl-, 57 dephospho-, 140 isobutyryl-, 94 methyl malonyl-, 93, 94, 431 radiolabeled, 140 succinyl-, 91, 93

SUBJECT INDEX Coenzyme B, 116–119, 123, 125–128, 236, 435, 436, 440–442, 451 density functional theory calculations, see Density functional theory calculations structure, 117, 436 Coenzyme B12 (and derivatives) (see also Adenosylcobalamin and Vitamin B12), 1–41 base-off, 31 base-off,His-on, 87 base-on, 31 biosynthesis, see Biosynthesis 2 0 -deoxy-, 4, 33 -dependent enzymes, 38, 83 derivatives as cofactor, 34–40 derivatives as intermediates in enzymes, 34–40 homo-, 4, 11, 14, 15 ‘‘inorganic’’ derivatives, 9, 13 molecular switch, see Molecular switch neo-, 8, 14, 15, 26 organometallic chemistry, 1–41 pseudo-, 8, 14, 15, 18, 26 reactivity, 24–34 redox chemistry, 18–24, 26 -rotaxane, 26, 27 spectroscopic studies, 13–16 structure, 4–18 Coenzyme (cofactor) F430, see F430 Coenzyme M, 57, 59, 67, 71–73, 118, 122, 123, 125, 126, 129, 236, 436 allyl-, 128 ethyl-, 125, 127, 128 list of methyl donors, 69 methyl transfer, see Methyl transfer methyl-, see Methyl-coenzyme M methyltransferase, see Methyltransferases reductase, see Reductases Cofactors computational studies, 417–452 F430, see F430 FeV, 370, 371 heme, see Heme(s) iron guanylylpyridinol, see Iron guanylylpyridinol cofactor iron-molybdenum, see FeMo cofactor trihydroxyphenylalanine quinone, see 2,4,5-Trihydroxyphenylalanine quinone

469 Combustion of fuels, 243 Complete active space second-order perturbation theory, 426 Compound I, 252, 285 Computational studies (of) ACS/CODH, 139, 144, 146 cofactors, 417–452 enzymes, 417–452 methods, 421–424 spectroscopic observables, 424–426 Conformational changes, 36, 59, 82, 91, 126, 156, 229, 247, 272, 275, 319, 320, 324, 326 dynamics, 15 Copper (different oxidation states) (in) 63,65 Cu, 334 alkyl complex, 349 at catalytic sites, 295–350 carbon bonds, see Copper-carbon bonds imidazole coordination, 298–349 MoCu CODH, see Molybdenum-copper carbon monoxide dehydrogenase Copper(I) (in), 295–350 alkyl complexes, 349 -arene p interactions, 306, 307, 344, 345 benzo[h]quinoline complex, 306 bleomycin, see Bleomycin carbon monoxide binding, 300–302, 311–315, 319–325, 328, 337, 341–343 copper amine oxidase, see Copper amine oxidase CusF chaperone, 344, 345 cyanide binding, 327–329, 331, 332, 343, 350 dioxygen binding, 300–302, 308, 310 ethylene receptor, see Copper-ethylene receptor inactivation of [Fe]-hydrogenase, 234, 235 nitrite reductase, see Nitrite reductase olefin interaction, 298, 345 receptors, 344–346 recognition sites, 344–346 stability constants, see Stability constants (synthetic) carbonyl complexes, 300–302, 305–308, 312–317, 323 -tryptophan interaction, 344, 345 Copper(II) (in), 295–350 alkyl complexes, 349 bleomycin, see Bleomycin copper amine oxidase, see Copper amine oxidase

Met. Ions Life Sci. 2009, 6, 461–496

470 [Copper(II) (in)] -nitric oxide, 330 nitrite reductase, see Nitrite reductase reduction, 331, 332 -superoxide complex, 310, 311 Copper(III), 349 Copper amine oxidase (from), 334–337 active site, see Active sites Arthrobacter globiformis, 334, 336 catalytic cycle, 335 Cu(I), 335–337 Cu(II), 335 cyanide inhibition, 335, 336 ping-pong mechanism, 335 structure, 334, 335 X-ray absorption spectroscopy studies, see X-ray absorption spectroscopy Copper-carbon bonds, 295–350 Copper chaperones CusF, 344, 345 Copper proteins bacterial copper chaperone CusF, 344, 345, 350 blue, 337–344 coupled binuclear, 298–308 green, 337–344 non-coupled binuclear, 308–317 purple, 337–344 Type III, 299 Copper-zinc superoxide dismutase, 330–332 active site, see Active sites bovine, 330, 331 cyanide binding, 331, 332, 376 cyanide probe, 368 electron transfer in, see Electron transfer hydrogen bonds, see Hydrogen bonds infrared studies, see Infrared spectroscopy mechanism, 331 Raman spectroscopy studies, see Raman spectroscopy structure, 331 Corallina officinalis, 369 Corphin coenzyme F430, see F430 origin of name, 117 Corrin(s) (see also individual names) alkylation, see Alkylation base-off, 22, 29 Co(I)-, 24–26, 35, 36 Co(II)-, 20–22, 24–26, 35

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX [Corrin(s) (see also individual names)] Co(III)-, 23, 26, 29, 32, 36 ‘‘complete’’, 25, 29 cyano-Co(III)-, 18 -dependent enzymes, 53–107 disproportionation, 21 ‘‘incomplete’’, 26, 29 methyl transfer, see Methyl transfer Corrinoid iron-sulfur protein, 56, 57, 59, 60, 73–76, 79, 80, 434, 443, 445, 446 Co(III), 136, 138–140 conformational change, 76 EPR studies, see EPR magnetic circular dichroism studies, see Magnetic circular dichroism Mo¨ssbauer spectroscopy studies, see Mo¨ssbauer spectroscopy structure, 75 Corrinoids (see also individual names) 1–41 adenosyl-, 34, 38–40 base-off, 34, 36, 55, 78 base-off/His-on, 34, 36, 38, 56 base-on, 34, 56 Co(I), 29, 58 Co(II), 5, 29, 33 Co(III), 5, 422 ‘‘complete’’, 4, 7–20, 29, 34 electron density, see Electron density epi-, 13 EPR studies, see EPR ‘‘incomplete’’, 5, 6, 13, 15, 21 methyl transfer, see Methyl transfer methyl-, 34, 74 Raman spectroscopy studies, see Raman spectroscopy structure, 4. 6 Corynebacterium diphtheriae, 248, 250, 252, 256–258, 276, 280, 283, 284 Crab horseshoe, 299, 306 Crosslinks cysteine-histidine, 303–305 cysteine-tyrosine, 337 histidine-tyrosine, 323, 324 Crystal structures (of) (see also X-ray crystal structures) adenosylcobalamin, 14 aldehyde oxidoreductase, 403 aquacobalamin, 9,10 carbonic anhydrase, 385, 387 coenzyme B12, 10

SUBJECT INDEX [Crystal structures (of) (see also X-ray crystal structures)] [Fe]-hydrogenase, 224, 228–230, 232 HybD protease, 164, 165 Hyp, 163, 164 manganese superoxide dismutase, 374, 376 methylcobalamin, 11, 14 methyl-coenzyme M reductase, 129 methylmalonyl-coenzyme A mutase, 90, 92, 431 [NiFe]-hydrogenase, 153–158, 162, 168. 172 [NiFeSe]-hydrogenase, 166 protocatechuate 3,4-dioxygenase, 380 ribonucleotide triphosphate reductase, 105 xanthine oxidoreductase, 398, 404–413 Curvularia inaequalis, 369 Cyanamide hydration, 385 Cyanate (in), 387 ACS/CODH, 135, 142 as inhibitor, 387 hydration, 385 thio-, see Thiocyanate Cyanidase, 366 Cyanide(s) (in), 297, 298, 350, 375 11 CN, 350 13 CN, 372 ACS/CODH, 135, 142–144 as inhibitor, 256, 269, 270, 274, 282–284, 337, 366, 372, 373, 381, 387, 388 as prebiotic substrate, 365, 366 as probe for transition metal sites, 366–368 biodegradation, 366 bridging, 328–330 copper amine oxidase inhibition, 335, 336 copper binding, see Copper(I) copper-zinc superoxide diamutase, see Copper-zinc superoxide dismutase cytochrome c oxidase, see Cytochrome c oxidase Fe(II), see Iron(II) Fe(III), see Iron(III) [FeFe]-hydrogenases, 179–208, 448 [Fe]-hydrogenases, 220, 221, 236 gold complex, 387 heme oxygenase, see Heme oxygenases heme-copper oxidases, see Heme-copper oxidases hydratase, see Hydratases iso-, see Isocyanide

471 [Cyanide(s) (in)] isotopically labeled, 331 managnese catalase, 377 manganese superoxide dismutase, 376, 377 monoxygenase, see Monoxygenases multi-copper oxidases, 339, 340 [NiFe]-hydrogenases, 158, 160–164, 171 non-heme iron enzymes, 381–383 oxidation, 372 oxygen evolving complex, 382, 383 protocatechuate 3,4-dioxygenase, 380, 382 reactions, 366 reduction, 372 -resistant respiratory systems, 365 superoxide reductase, 381 toxicity, see Toxicity vanadium enzymes, 372, 373 xanthine oxidoreductase, 397 zinc enzymes, 387, 388 zinc hydrolase, 388 Cyanidium caldarium, 246 Cyanobacteria (see also individual names), 181, 246, 283, 284, 373 Cyanocobalamin (see also Vitamin B12), 87 Co(II), 23 Co(III), 3, 23 Cyanohydrin, 365 Cyclase guanylate, see Guanylate cyclase Cyclic voltammetry studies of B12 derivatives, 22 Cyclodextrins, 27 Cysteine -histidine crosslink, see Crosslinks homo-, see Homocysteine seleno-, see Selenocysteine Cytochrome aa3, 327–329 Cytochromes ba3, 327–329 Cytochrome bo3, 318–321, 324 EPR studies, see EPR oxidase, 318, 320, 321, 324, 326, 327 Cytochrome c, 165, 166 Cytochrome c oxidase, 244, 317, 319–322, 343, 365, 366 active site, see Active sites bovine (heart), 318, 320, 326, 329 cyanide binding, 327–329 electron transfer in, see Electron transfer Fe(II)-carbon monoxide, 318 mammalian, 325, 330

Met. Ions Life Sci. 2009, 6, 461–496

472

SUBJECT INDEX

[Cytochrome c oxidase] mutagenesis, 323 nitric oxide inhibition, 329, 330 photolysis, 318, 324, 326 proton transfer in, see Proton transfer structure, 323 Cytochrome P450 cam, 253–255 reductase, see Reductases Cytotoxicity of bleomycin, 349

D Decarboxylase uroporphyrinogen, 68 Deformylase peptide, 379 Degradation of aromatic molecules, 378, 380 Dehalogenation reductive, 34, 35, 60, 77, 78, 120 Dehydrases diol, see Diol dehydrase propanediol, 84 Dehydrogenases alcohol, see Alcohol dehydrogenase carbon monoxide, see Carbon monoxide dehydrogenase F420-dependent methylenetetrahydromethanopterin, 222 H2-forming methylenetetrahydromethanopterin, see [Fe]-hydrogenases xanthine, 404, 405, 410 O-Demethylases, 59, 76, 77 vanillate, 77 Denitrification bacterial, 342 copper enzymes in, 340–344 Density functional theory calculations, 419–452 acetyl-coenzyme A synthase, 445, 446 cobalamins, 86, 425, 429, 431, 434, 435 cobinamides, 425 coenzyme B, 123 F430, 125, 127, 437 [FeFe]-hydrogenases, 189, 192, 193, 198, 199, 447, 449

Met. Ions Life Sci. 2009, 6, 461–496

[Density functional theory calculations] H-cluster of [FeFe]-hydrogenases, 449, 450 heme-copper oxidases, 324 methyl-coenzyme M reductase, 120, 440–442 methyl-coenzyme M, 123 MoCu CODH, 333, 334 [NiFe]-hydrogenases, 170, 447 peptidylglycine a-hydroxylating monooxygenase, 310 time-dependent, 426, 429, 434, 438 truncated models, 434, 435, 438 5 0 -Deoxyadenosylcobalamin, see Coenzyme B12 Deoxyhemocyanin, 299 active site, 300 Deoxyribonucleic acid, see DNA Deprotonation constants, see Acidity constants Desulfitobacterium dehalogenans, 78 frappieri, 78 Desulfomicrobium baculatum, 156, 166 Desulfovibrio desulfuricans, 152, 156, 166, 168, 181–192 fructosovorans, 156, 165–169 gigas, 153, 155, 156, 168, 187, 398, 403, 411 vulgaris, 153, 156, 168, 189, 190 Detoxification of chlorinated compounds, 34 environmental pollutants, 340 reactive oxygen species, 374 Deuterium 2 H NMR, see NMR CD3-Ni(II)F430M, 120 hydrogen exchange, 168, 269–272, 275 isotope effects, 92 label, 96, 101, 106 Diazotrophs, 370 Dielectric constant, 434 DFT, see Density functional theory calculations Dihydrogen (see also Hydrogen), 172, 221, 225, 236, 237 acidity constant, see Acidity constants activation, 221, 222 catalytic cycle, 197 formation, 225

SUBJECT INDEX [Dihydrogen (see also Hydrogen)] heterolytic cleavage, 168, 172, 191, 235 production, 370, 371 3,4-Dihydroxybenzoate, see Protocatechuate 2,4-Dihydroxypteridine, see Lumazine 2,6-Dihydroxypurine, see Xanthine 4,6-Dihydroxypyrazolo[3,4-d]-pyrimidine, see Alloxanthine 5,6-Dimethylbenzimidazole, 4, 8, 12–16, 19, 20, 25, 29, 30, 32, 33, 36, 37, 54–56, 62, 79, 87, 90, 424, 428, 429 15 N-labeled, 96 -cobamide, see Cobalamins -off/His-on, 87, 90, 103 -on, 96, 101, 106 structure, 428 Dimethylformamide, 29 2,6-Dimethylphenyl isocyanide (in) dopamine b-monooxygenase, 315–317 peptidylglycine a-hydroxylating monooxygenase, 315–317 Dinitrogen (see also Nitrogen) fixation, see Nitrogen fixation reduction, 370, 373 gem-Diol, 383 Diol dehydrase, 11, 38, 84, 85, 95–101 conformational change, 100 hydrogen bonds, see Hydrogen bonds mechanism for hydroxyl migration, 97–100 structure, 96 Dioxygen (see also Oxygen) (in), 153. 169, 225, 244, 245, 330, 335 17 O2, 170, 171 activation, 247, 251–259, 265, 275, 276, 285, 314, 325, 378 Cu(I) binding, see Copper(I) heme-copper oxidases, 318 hemocyanin, 298 O–O bond cleavage, 255, 256, 267, 268, 379 production, 373–375, 381 rate constants, see Rate constants reduction, 299, 317, 324, 330, 335, 337, 339, 340, 379 transfer, 319 xanthine oxidoreductase, 396 Dioxygenases catechol, see Catechol dioxygenases protocatechuate, see Protocatechuate dioxygenase Rieske, 378

473 Diseases Lou Gehrig’s, see Amyotrophic lateral sclerosis Disulfide, 105, 451 bonds, 104, 441, 442 bridge, 228 diironhexacarbonyl-, 195, 196 hetero-, 116, 123 Dithiol(ate) (see also Thiols), 187, 195, 197, 204, 205 non-protein, 191, 192, 195 propane, 188, 191, 192, 196, 197, 204 Dithionite (in) ACS/CODH, 137, 142, 144 [FeFe]-hydrogenase, 448 sodium, 188, 282 Dithiothreitol, 81, 228 Diuretics, 387 DMB, see 5,6-Dimethylbenzimidazole DNA damage, 41, 347–349 hydrolysis, 386 Dopamine, 308 Dopamine b-monooxygenase, 308–312, 314–316 active site, see Active sites carbon monoxide reactions, see Carbon monoxide Fourier transform infrared spectroscopy studies, see Fourier transform infrared spectroscopy infrared studies, see Infrared spactroscopy X-ray absorption spectroscopy studies, see X-ray absorption spectroscopy Drugs (see also individual names), 404 antiinflammatory, 378

E EDTA, see Ethylenediamine-N,N,N 0 ,N 0 tetraacetate Electrochemistry of B12 derivatives, 26, 27 Electrodes normal hydrogen, 117, 124, 339, 372 saturated calomel, 22, 26, 29 standard hydrogen, 75, 429 Electron density (in), 366 blue copper proteins, 338 carbon monoxide dehydrogenase, 141, 142

Met. Ions Life Sci. 2009, 6, 461–496

474 [Electron density (in)] cobalamins, 87 corrinoids, 70, 71 Cu(I) complex, 306 [FeFe]-hydrogenases, 188, 191 [Fe]-hydrogenase, 233 haloperoxidases, 372 heme oxygenase, 260, 261 iron-guanylylpyridinol cofactor, 230–232 superoxide reductase, 382 Electron nuclear double resonance spectroscopy (studies of) 13 C, 101 1 H, 254 2 H, 101 ACS/CODH, 139, 142 continuous wave, 123 cryogenic, 252 [FeFe]-hydrogenases, 192, 194 H217O, 171 heme oxygenase, 253, 254, 259, 260 methyl-coenzyme M reductase, 121, 440 [NiFe]-hydrogenases, 170, 171 pulse, 123 xanthine oxidoreductase, 397, 400–403, 405, 408 Electron paramagnetic resonance, see EPR Electron spin echo envelope modulation, 146 [FeFe]-hydrogenases, 192–194 xanthine oxidoreductase, 402, 405 Electron spin resonance, see EPR Electron transfer (in) ACS/CODH, 135, 444 B12 derivatives, 19, 2–23, 26, 35 blue copper proteins, 337–339, 342, 343 cobalamins, 77 copper amine oxidase, 335 Cu-Zn superoxide dismutase, 331 cytochrome c oxidase, 324 [FeFe]-hydrogenases, 184 heme oxygenases, 247 heme-copper oxidases, 318, 324–327 hemocyanins, 299 methionine synthase, 61 [NiFe]-hydrogenases, 165, 166 oxygen evolving complex, 373, 374 peptidylglycine a-hydroxylating monooxygenase, 313 respiratory chains, 366 ribonucleotide triphosphate reductase, 104–106

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX [Electron transfer (in)] superoxide reductase, 381, 382 xanthine oxidoreductase, 403 Electrostatic interaction in heme oxygenases, 279, 280 trafficking, 331 ENDOR, see Electron nuclear double resonance spectroscopy Enzymes (see also individual names) catalyzing carbon skeleton rearrangements, 86–96 cobalamin-dependent, 53–107 coenzyme B12-dependent, 38, 83 computational studies, 417–452 corrinoid-dependent, 53–107 manganese, 373–377 molybdenum, 395–413 non-heme iron, 377–388 tungsten, 398 vanadium, 368–373 zinc, 383–388 EPR (studies of), 367 17 O, 171, 403 2D, 15 acetyl-coenzyme A synthase, 445 ACS/CODH, 137, 139, 142–144, 146, 443 blue copper proteins, 338 Co(II)-corrinoids, 15 cob(II)inamide, 424, 425 cobalamins, 424, 425, 429 continuous wave, 120, 121 corrinoid iron-sulfur protein, 65, 74 cytochrome bo3, 326 diol dehydrase, 96, 97 Fe(III)-OOH, 252, 253, 259 [FeFe]-hydrogenases, 192, 193, 197 glutamate mutase, 89 manganese catalase, 377 methionine synthase, 65 methyl-coenzyme M reductase, 120–122, 126, 438, 440 methylmalonyl-coenzyme A mutase, 92 MoCu CODH model complex, 334 [NiFe]-hydrogenases, 153, 154, 168, 170, 172 nitrogenases, 371 oxygen evolving complex, 383 pulsed, 101, 402 rapid freeze quench, 104 single crystal, 172 superoxide reductase, 381

SUBJECT INDEX [EPR (studies of)] ‘‘very rapid’’ signal, 399, 408 xanthine oxidoreductase, 396–399, 402, 408 Equilibrium constants (see also Acidity constants and Stability constants) ACS/CODH, 139, 140 carbon monoxide binding to Cu(I) in heme-copper oxidases, 328 dioxygen binding to Cu(I) in hemocyanin, 300 Escherichia coli, 35 bo3 oxidase, 318, 320, 321, 324, 326 copper homeostasis, 344 ethanolamine ammonia lyase, 100 [FeFe]-hydrogenase, 201, 206 [Fe]-hydrogenase, 227, 228 heme oxygenase, 251 iron superoxide dismutase, 376 isobutyryl-coenzyme A mutase genes, 94 metabolism, see Metabolism methionine synthase, 60–67 [NiFe]-hydrogenases, 161, 162, 164 NikR, 223 ornithine 4,5-aminomutase, 103 ESEEM, see Electron spin echo envelope modulation ESR, see EPR Ethanolamine, 100, 101 Ethanolamine ammonia lyase, 11, 38, 85, 95, 96, 100, 102, 201 reaction mechanism, 100 Ethylene perchloro-, 77, 78 receptor, 345 tetrachloro-, 35, 78 Ethylenediamine-N,N,N 0 ,N 0 -tetraacetate, 235 Eubacteria (see also individual names), 84, 332, 442 anaerobic, 57 Eukaryotes (or eukaryotic) (see also individual names), 56, 181, 184, 334, 374 methionine synthase, 60 European Synchrotron Radiation Facility, 157, 158 Evolution of life, 365 EXAFS, see Extended absorption fine structure spectroscopy

475 Extended absorption fine structure spectroscopy (studies of) heme-copper oxidases, 321 K-edge, 372 MtaA, 71 [NiFe]-hydrogenases, 156 [NiFeSe]-hydrogenases, 157 peptidylglycine a-hydroxylating monooxygenase, 312, 313, 316

F F430 (see also Methyl-coenzyme M reductase), 116, 419, 435–442, 446 absorption spectroscopy, see Absorption spectroscopy active site, see Active sites catalytic cycle, 117, 125 density functional theory calculations, see Density functional theory calculations magnetic circular dichroism studies, see Magnetic circular dichroism methylation, see Methylation methyl-Ni(III), 122, 123, 125 model, 438, 439 Ni(I), 117, 120, 121, 125, 126, 128, 437–439 Ni(II), 117, 125, 437–439, 441 Ni(III), 120–125, 437–439, 451 pentamethyl ester, see F430M redox potential, see Redox potentials Siegbahn mechanism, 128 stability constants, 119 structure, 117–119, 436 3-sulfonatopropyl-Ni(III)-, 120–122 F430M methyl-, 120 Ni(I), 117, 119 Ni(II), 117, 119, 120 Ni(III), 117, 120 redox couples, see Redox potentials stability constants, 119 Facial triad motif 2-His-1-carboxylate, 378–383 structure, 378 Factor A, 16, 18 structure, 4, 9, 14 Fatty acids, 378 Febuxostat, see FYX-051

Met. Ions Life Sci. 2009, 6, 461–496

476 [FeFe]-hydrogenases (from), 152, 179–208, 220, 221, 234, 447–452 active site, see Active sites amino acid sequences, 223 biosynthesis, see Biosynthesis carbon monoxide in, see Carbon monoxide Chlamydomonas reinhardtii, 183, 200, 201 Clostridium pasteurianum, 181–195, 207 cyanide in, 179–208, 448 density functional theory calculations, see Density functional theory calculations Desolfovibrio vulgaris, 189–194 Desulfovibrio desulfuricans, 181, 182, 184–195, 197 electron density, see Electron density electron transfer in, see Electron transfer ENDOR studies, see ENDOR spectroscopy EPR studies, see EPR ESEEM studies, see Electron spin echo envelope modulation Fe(I), 448 2Fe subcluster, 187–190, 192–199, 204, 448–450 Fourier transform infrared spectroscopy studies, see Fourier transform infrared spectroscopy genes, 200–202 H-cluster, 182–198, 199, 447–449, 451 H-cluster biosynthesis, 199–207 H-cluster models, 195–199, 449, 450 hyperfine sublevel correlation spectroscopy studies, see Hyperfine sublevel correlation spectroscopy studies of infrared studies, see Infrared spactroscopy inhibition by carbon monoxide, 190–193, 448 Mo¨ssbauer spectroscopy studies, see Mo¨ssbauer spectroscopy occurrence, 181 oxidation states of the H-cluster, 189, 192–195, 197, 198 photochemical cleavage, 190, 191, 193 ping-pong catalytic mechanism, 222 sequence alignments, 186 spectroscopic studies, 192–195 structure, 181–192, 447, 448 Thermotoga maritima, 186

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX [[FeFe]-hydrogenases (from)] X-ray diffraction spectroscopy studies, see X-ray diffraction spectroscopy [Fe]-hydrogenases (from), 152, 181, 219–237 active site, see Active sites amino acid sequences, 223 apo-, 224, 227–229, 232 biosynthesis, see Biosynthesis carbon monoxide in, 219–237 carbon monoxide in, see Carbon monoxide catalytic mechanism, 222, 235–237 conformational change, 229 crystal structure, see Crystal structures cyanide in, 220, 221, 233 electron density, see Electron density extrinsic ligands, 232–234 Fourier transform infrared spectroscopy studies, see Fourier transform infrared spectroscopy genes, 223 holo-, 227–233 inactivation by Cu(I), 234, 235 inactivation by superoxide, 234, 235 infrared studies, see Infrared spactroscopy inhibition, 220, 232–234, 236 intrinsic ligands, 231, 232 iron ligands, 229–235 Methanocaldococcus jannaschii, 227–230 Methanopyrus kandleri, 222, 223, 227–229 Methanothermobacter marburgensis, 223, 227, 233, 234 Mo¨ssbauer spectroscopy studies, see Mo¨ssbauer spectroscopy NMR studies, see NMR physiology, 222, 223 protein fold, 228 structure, 227–230 X-ray diffraction spectroscopy studies, see X-ray diffraction spectroscopy FeMo cofactor, 370 biosynthesis, see Biosynthesis Fenton chemistry, 161 [Fe]-only hydrogenases, see [FeFe]hydrogenases Ferredoxins (see also individual names), 75, 80, 162, 182–184 carbon monoxide dehydrogenase, 445 Ferritin, 245 [Fe2S2] cluster in [FeFe]-hydrogenases, 182–184, 205 xanthine oxidoreductase, 403

SUBJECT INDEX [Fe3S4] cluster in ACS/CODH, 144, 155, 156, 165, 169, 170 corrinoid-dependent enzymes, 78 [Fe4S4] cluster (in) ACS/CODH, 134, 136–139, 141, 146, 442–450, 452 corrinoid-dependent enzymes, 73, 75, 78, 80 [FeFe]-hydrogenases, 182–184, 187–190, 192–194, 198, 204 [Fe4S4]1+, 137, 138, 145, 146, 193 [NiFe]-hydrogenases, 155, 156, 162, 165 nitrogenases, 370 Flavodoxins, 62, 65–67, 79–82, 154, 155 as chaperone, 81 reductase, see Reductases Fluorescence resonance energy transfer heme oxygenase, 247 Force field calculations scaled quantum mechanical, 425 self-consistent field, see Self-consistent field method Formaldehyde, 373 Formate, 60, 72, 76 Formation constants, see Equilibrium constants and Stability constants Fosfomycin, 82–84 14 C-labeled, 83 biosynthesis, see Biosynthesis methyl transfer, see Methyl transfer Fourier transform infrared spectroscopy (studies of) dopamine b-monooxygenase, 314, 315 [FeFe]-hydrogenases, 188, 194, 197 [Fe]-hydrogenases, 232, 233 heme-copper oxidases, 319, 320, 326, 329 iron guanylylpyridinol cofactor, 224–226 [NiFe]-hydrogenases, 158, 159, 172, 187 step-scan, 319, 320, 324 FRET, see Fluorescence resonance energy transfer Fructose-1,6-diphosphate aldolase, 383 FTIR, see Fourier transform infrared spectroscopy Fungi (see also individual names) nitrogen source, 366 tyrosinase, see Tyrosinase

477 [Fungi (see also individual names)] vanadium chloroperoxidase, 369 vanadium haloperoxidase, 372 FYX-051 as inhibitor, 405. 406 complex with xanthine oxidase, 405, 406

G Galactose oxidase, 337 active site, see Active sites inhibition by cyanide, 337 nuclear magnetic relaxation dispersion measurements, 337 Genes clusters, 201, 202 hmd, 223, 227 hyd, 201, 202 narF, 185 nif, 200 nikR, 223 Genome of Methanosarcina acetivorans, 67, 82 Globins hemo-, see Hemoglobin hydride complex, 261 myo-, see Myoglobin Glucose fermentation, 59, 72 6-phosphatase, 369 Glutamate mutase, 11, 62, 86–92, 431, 432 active site, see Active sites radical formation, 88 reaction mechanism, 88 Glutamate synthase NMR studies, see NMR Glycine radical, see Radicals Glycolaldehyde, 84 Gold cyanide complex, 387 G-protein, 345, 346 GTP deoxy-, 104, 106 GTPases, 93, 164, 204 Guaiacol, 253 Guanidinium group, 101 Guanosine triphosphate, see GTP Guanylate cyclase, 244

Met. Ions Life Sci. 2009, 6, 461–496

478

SUBJECT INDEX

H Haloperoxidases electron density, see Electron density vanadium, 368–370, 372 Hartree-Fock theory, 422 Helix pomatia, 302, 308 Heme(s) (see also individual names) 13 C-labeled, 260 a-meso-hydroxy-, 246–248, 256–259, 276, 277, 284, 285, 318–325 a3, 244, 318, 319, 324, 325, 329 active site, see Active sites as cofactor, 241–285 as substrate, 241–285 breakdown, 246–248, 268–276, 284 -copper oxidases, see Heme-copper oxidases dioxygen binding, 276 hydroxide complexes, 260–262 hydroxylation, 247, 250–254, 259–268, 275–284 iron release, 245, 248 model complexes, 259, 260 non-heme iron enzymes, see Non-heme iron enzymes oxidation, 246, 247, 251, 279–284 oxygenase, see Heme oxygenases regioselectivity of hydroxylation, 275–284 regioselectivity of oxidation, 279–284 verdo-, 246, 247, 252, 256–259, 276 Heme-copper oxidases (see also individual names), 317–330 active site, see Active sites bacterial, 330 carbon monoxide in, see Carbon monoxide catalytic cycle, 325 cyanide binding, 319–325 dioxygen binding, 318 electron transfer in, see Electron transfer EXAFS studies, see Extended absorption fine structure spectroscopy Fourier transform infrared spectroscopy studies, see Fourier transform infrared spectroscopy hydrogen bonds, see Hydrogen bonds infrared studies, see Infrared spectroscopy linkage isomers, 328, 329 mixed-valent, 324–327 photolysis, 319, 324, 325 proton transfer, see Proton transfer

Met. Ions Life Sci. 2009, 6, 461–496

[Heme-copper oxidases (see also individual names)] Raman spectroscopy studies, see Raman spectroscopy structure, 318, 321 synthetic models, 318, 329 Heme oxygenases (from), 243, 246 -1, 245–259, 273, 274, 276, 277, 279, 280, 284 -2, 245, 249, 251, 252 -3, 245 an ubiquitous enzyme, 245–247 apo-, 249 azide complex, 263–267, 269–274, 276, 277 azide inhibition, 269, 270, 274 bacterial, 247, 248, 250, 252, 258, 264, 269, 282 carbon monoxide complex, 271, 274, 275 carbon monoxide inhibition, 269, 274, 278, 285 catalytic cycle, 247, 248, 251, 252, 258, 268, 269, 274–276 conformational changes, 271, 273, 275 Corynebacterium diphtheriae, 248, 250, 252, 256–258, 276, 280, 283, 284 cyanide complex, 262–265, 267, 269–274, 284 cyanide inhibition, 256, 269, 270, 274, 282–284 density functional theory calculations, see Density functional theory calculations dynamics, 268–276 electron density, see Electron density electron transfer in, see Electron transfer ENDOR studies, see ENDOR spectroscopy ferryl formation, 259–268 heme-heme, 244 heteronuclear single quantum coherence spectroscopy studies, see Heteronuclear single quantum coherence spectroscopy holo-, 249 homolytic dioxygen cleavage, see Dioxygen human, 148, 249, 251, 253–256, 264, 274, 276, 284 hydrogen bonding network, see Hydrogen bonds isoforms, 245

SUBJECT INDEX [Heme oxygenases (from)] mammalian, 247, 249, 250, 255, 256 mutant, 253–256, 272–274, 279–282 Neisseria meningitidis, 248, 250, 255–258, 262, 274, 278, 280–284 PDB codes, 249 plant, 246, 247 Pseudomonas aeruginosa, 248, 250, 252, 256–258, 260–284 rat, 248, 250, 252, 253, 255, 264, 276, 277, 279, 280, 284 reactivity, 274–276 structure, 248–251, 253, 255, 256, 270, 276, 279, 282 structure-function relationship, 253 wild-type, 253, 254 X-ray diffraction spectroscopy studies, see X-ray diffraction spectroscopy Heme proteins (see also individual names) Fe(III), 367 Hemocyanin(s), 297–308 active site, see Active sites arthropodal, see Arthropods carbon monoxy-, 304–308 deoxy-, see Deoxyhemocyanin electron transfer in, see Electron transfer molluscan, see Molluscs oxy-, see Oxyhemocyanin structures, 299, 304 Hemoglobin, 299, 302, 367 carbon monoxide binding, 244, 298 carboxy-, 244 human, 301 Heteronuclear single quantum coherence spectroscopy (studies of) 1 H,15N, 275 heme oxygenase, 265, 274, 275 High-performance liquid chromatography, 223 UV, 28 Histidine 15 N-, 56 -coordinating cluster, 184 -cysteine crosslink, see Crosslinks His-on, 34, 36–38, 56, 65, 66, 70, 87, 90, 102, 103 His-off, 65, 66 kinase, 284 stacking, see Stacking Hodgkin’s lymphoma, 347

479 Homeostasis of (see also Metabolism) copper, 344 Homocysteine (in) corrinoid methyltransferases, 58 methionine synthase, 60, 61, 63–66, 432–434 structure, 60 Homocitrate in FeMo cofactor, 370 Hormones, 245 HPLC, see High-performance liquid chromatography Human carbonic anhydrase, see Carbonic anhydrase heme oxygenase, see Heme oxygenase hemoglobin, see Hemoglobin metabolism, see Metabolism methionine synthase, 60, 80 methylmalonyl-coenzyme A mutase, 92 myoglobin, see Myoglobin HSQC, see Heteronuclear single quantum coherence spectroscopy Hydratases cyanide, 366 nitrile, 379 Hydrazine, 372 formation, 371 Hydride bridge, 236 transfer, 57, 221 Hydrogen (see also Dihydrogen) 1 H ENDOR, see ENDOR 1 H NMR, see NMR 1 H,15N HSQC, see Heteronuclear single quantum coherence spectroscopy 2 H, see Deuterium 3 H, see Tritium bond, see Hydrogen bond molecular, see Dihydrogen oxidation, 165, 171, 181, 182, 190, 191, 198 production, 153, 173 reducing power, 152 transfer, see Transfer Hydrogenases, 57, 80, 82 classes, 152 [FeFe], see [FeFe]-hydrogenases [Fe], see [Fe]-hydrogenases [NiFe], see [NiFe]-hydrogenases [NiFeSe], see [NiFeSe]-hydrogenases regulatory, 161

Met. Ions Life Sci. 2009, 6, 461–496

480 Hydrogen bonds (in/to) ACS/CODH, 143 carbonic anhydrase, 387 cobalamins, 15, 62, 67, 68, 87, 90, 429 Cu-Zn superoxide dismutase, 331 diol dehydrase, 98–100 heme oxygenase, 249, 250, 253–256, 258, 264–267, 272–275, 279–281, 285 heme-copper oxidases, 324 manganese catalases, 376 methyl-coenzyme M reductase, 441 [NiFe]-hydrogenases, 158, 166 nitrite reductase, 341 peptidylglycine a-hydroxylating monooxygenase, 313 phosphatases, 369 protocatechuate 3,4-dioxygenase, 382 xanthine oxidoreductase, 400, 401, 403 Hydrogen peroxide, 252, 253, 330, 331, 335, 337, 368, 372, 374, 376, 381 disproportionation, 375 Hydrolase zinc, see Zinc hydrolase Hydrolysis ACS/CODH, 141 5 0 -ATP, 75, 82, 373 carboxylic acid esters, 385 DNA, see DNA RNA, see RNA sulfonic acid esters, 385 Hydroperoxide(s) (in) (see also Peroxides) acyl-, 252 alkyl-, 252 bridging, 169–171 ferric, see Iron(III) formation, 251, 252 [NiFe]-hydrogenases, 168–171 Hydrothermal vents, 199, 222 Hydroxide(s), 143 bridging, 137, 142, 188 complex with heme oxygenase, 261, 262, 274 Hydroxocobalamin, 74 base-on, 75 Co(III), 3, 19, 20 structure, 4 Hydroxo group, 135 bridged, 168, 169 Hydroxylase(s) (see also individual names) molybdenum-containing, 395–413

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX Hydroxyl group migration, 96–101, 103 transfer, see Transfer 4-Hydroxypyrazolo[3,4-d]-pyrimidine, see Allopurinol Hyperfine sublevel correlation spectroscopy studies of [FeFe]-hydrogenases, 194 methyl-coenzyme M reductase, 121, 123 [NiFe]-hydrogenases, 172 Hyperuricemia, 404 Hypohalous acids, 368 Hypoxia, 244 HYSCORE, see Hyperfine sublevel correlation spectroscopy studies of

I Imidazole (and moieties) (in) (see also Histidine), 12, 25, 33, 37, 56 ACS/CODH, 136 bridge, 331, 332 copper coordination, 298–349 1,2-dimethyl-, 316 5,6-dimethylbenz-, see 5,6Dimethylbenzimidazole 1-methyl-, 314 Ni(II) complex, 119 Immune response, 329 Infection bacterial, 246 Infrared spectroscopy (studies of) ACS/CODH, 139, 141 Cu(I) complex, 305, 316 Cu,Zn superoxide dismutase, 368 dopamine b-monooxygenase, 311 [FeFe]-hydrogenases, 189, 192–194, 197 [Fe]-hydrogenase, 229, 231, 233 Fourier transform, see Fourier transform infrared spectroscopy heme-copper oxidases, 319, 320, 322, 324, 328 iron guanylylpyridinol cofactor, 224, 226, 231 [NiFe]-hydrogenases, 172 peptidylglycine a-hydroxylating monooxygenase, 311 time-resolved, 322

SUBJECT INDEX Inhibition (of/by) azide, 269, 270, 274, 376, 377 bromoperoxidase, 372 3-bromopropane sulfonate, 120 carbonic anhydrase, 387 carbon monoxide, 190–193, 244, 269, 274, 278, 285, 311–316, 319–325, 336, 341, 342, 448 carbon monoxide dehydrogenases, see Carbon monoxide dehydrogenases cyanate, 387 cyanide, see Cyanide [FeFe]-hydrogenases, 190–193, 448 [Fe]-hydrogenases, 220, 232–234, 236 galactose oxidase, 337 heme oxygenases, see Heme oxygenases nitric oxide, 329, 330 methyl-coenzyme M reductase, see Methyl-coenzyme M reductase nitrite reductase, 341 respiration, 327 xanthine oxidoreductase, 404–406 Inhibition constants, 120,387 Interdependency magnesium–manganese, 374 Iodide, 369 oxidation, 372 propyl, 73 Iodoacetamide as scavenger, 92 Iodoperoxidase vanadium, 369 Ipomoea, 303 IR, see Infrared spectroscopy Iron (different oxidation states) (in) 57 Fe, 225, 232, 444, 445, 448 [NiFe]-hydrogenases, see [NiFe]hydrogenases oxidation states in [FeFe]-hydrogenases, 193, 197, 198 recycling, 246 Iron(0), 225, 232, 236 Iron(II) (in) ACS/CODH, 137, 138, 444 as substrate, 339 bleomycin, see Bleomycin catechol dioxygenase, 380 cyanide binding, 328–330, 365, 367 Fe(II)-CO, 318, 320–323 Fe(II)-O2, 247, 251–253, 276 -heme, 326

481 [Iron(II) (in)] 2-His-1-carboxylate coordination, 378–382 hydride, 236 oxidation, 340 oxygen evolving complex, 381 superoxide reductase, 381, 382 Iron(III) (in), 339, 340 ACS/CODH, 137 cyanide binding, 327–330, 365, 367 Fe(III)-, 267, 268 Fe(III)-OO–, 252–256, 274, 285, 378, 381 Fe(III)-OOH, 247, 248, 250–256, 259, 260, 262, 265, 267, 268, 276–278, 280–282, 285, 347, 348, 378, 382 -heme, 326 oxygen evolving complex, 381 protocatechuate 3,4-dioxygenase, 382 redox potential, see Redox potentials reduction, 247 superoxide reductase, 381, 382 Iron(IV) Fe(IV)=O, 252, 378 hydride, 236 Iron(V) Fe(V)=O, 247, 248, 378 Iron guanylylpyridinol cofactor, 221–232, 236 57 Fe-labeled, 225, 232 biosynthesis, see Biosynthesis electron density, see Electron density Fourier transform infrared spectroscopy studies, see Fourier transform infrared spectroscopy infrared studies, see Infrared spectroscopy isolation, 223, 224 matrix-assisted laser desorption/ionization time-of-flight mass spectrometry studies, 224, 225 Mo¨ssbauer spectroscopy studies, see Mo¨ssbauer spectroscopy stability, 225–227 structure, 224, 225 X-ray absorption spectroscopy studies, see X-ray absorption spectroscopy X-ray diffraction spectroscopy studies, see X-ray diffraction spectroscopy Iron-molybdenum cofactor, see FeMo cofactor

Met. Ions Life Sci. 2009, 6, 461–496

482

SUBJECT INDEX

Iron-sulfur proteins (see also individual names) corrinoid, see Corrinoid iron-sulfur proteins Irradiation UV, 226 Isobutyryl coenzyme A mutase, 86, 94 Isocyanides, 298, 315, 333, 345, 350, 372 2,6-dimethylphenyl-, see 2,6Dimethylphenyl isocyanide binding in myoglobin, 317 isopropal, 316 tert-butyl, 333 Isomerases phosphomannose, 383 SlyD, see Peptidyl prolyl cis/trans isomerase Isotope(s) (see also individual elements and compounds) exchange, 168, 269–272, 275 Isotope effects, 91, 158, 267 kinetic, 92, 96, 97

K Kinases histidine, 284 Klebsiella oxytoca, 100

L Laccase, 339, 340 b-Lactamase zinc, 386 Lactobacillus leichmannii, 104, 105 plantarum, 375 Laminaria digitata, 369 Leucine aminopeptidase, 386 Leukotrienes, 378 Ligases zinc, 383 Lignin degradation, 340 Limpet keyhole, 305 Limulus polyphemus, 299, 301–303, 306 Lipoic acid biosynthesis, see Biosynthesis

Met. Ions Life Sci. 2009, 6, 461–496

Lipoxins, 378 Lipoxygenases, 378 Loligo pealii, 305 Lumazine complex with xanthine oxidoreductase, 406–409 Luminescence carbon monoxyhemocyanin, 307 Lyases (see also individual names) ammonia aspartate, 202 ethanolamine ammonia, see Ethanolamine ammonia lyase zinc in, 383 Lysine aminomutases 2,3-, 95, 203, 205 5,6-, 101–103

M Macroconstants, see Acidity constants and Stability constants Magnesium(II) (in) alkaline phosphatase, 388 dimethyl, 120 interdependency with manganese, 374 [NiFe]-hydrogenases, 156, 166 Magnetic circular dichroism (studies of) cobalamins, 426, 429, 431 corrinoid iron-sulfur protein, 74 F430, 439 methyl-coenzyme M reductase, 437 nitrogenases, 371 superoxide reductase, 381 MALDI-TOF, see Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry Mammalian carbon monoxide in, 245 carbonic anhydrase, 385 cytochrome c oxidase, see Cytochrome c oxidase heme oxygenase, see Heme oxygenase iron level, 246 methionine synthase, 60, 81 methylmalonyl-coenzyme A mutase, 84 myoglobin, 251 skin, 300 tyrosinase, see Tyrosinase xanthine oxidoreductase, 405

SUBJECT INDEX Manganese (different oxidation states) (in) biology, 373, 374 interdependency with magnesium, 374 Manganese(II) (in) catechol dioxygenase, 380 Manganese catalases (from), 375, 376 active site, see Active sites catalytic cycle, 376 cyanide in, 377 hydrogen bonds, see Hydrogen bonds inhibition by azide, 377 Lactobacillus plantarum, 375 structure, 375 Thermus thermophilus, 375 Manganese enzymes (see also individual names), 373–377 Manganese superoxide dismutase (from), 374–376 azide in, see Azide crystal structure, see Crystal structures cyanide interaction, 376, 377 Thermus thermophilus, 376 types, 374 Mass spectrometry, 28, 123 MALDI-TOF, see Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry iron guanylylpyridinol cofactor, 224, 225 Matrix metalloproteinases, 386 MCD, see Magnetic circular dichroism Megathura crenulata, 305 Melanin biosynthesis, see Biosynthesis 2-Mercaptoethanesulfonate, see Coenzyme M Mercaptoethanol, 223, 224, 226, 227, 232 7-Mercaptoheptanoylthreonine, see Coenzyme B Mercury(II), 32 Metabolism (of) (see also Homeostasis) amino acids, 35 anaerobic, 204 carbohydrate, 201 carbon monoxide, 444 energy, 200, 221 Escherichia coli, 161 human, 34, 35, 38, 56 hydrogen, 447 microbial, 204

483 [Metabolism (of) (see also Homeostasis)] one-carbon, 35 roles of corrinoid-dependent methyltransferases, 56–60 Metaphosphate, 369 Methane, 35, 73, 119, 120, 127 chloro-, 78 dichloro-, 119 formation, see Methanogenesis monooxygenase, see Monooxygenases oxidation, 243 tetrachloro-, 120 Methane monooxygenase, 378 Methanobacterium thermoautotrophicum, 58, 72, 80 Methanobrevibacterium smithii, 223 Methanocaldococcus jannaschii, 223, 227–230 Methanococcus jannaschii, 164 maripaludis, 223 voltae, 155 Methanococcoides sp., 67 Methanogenesis, 35, 57–59, 72, 73, 116, 123, 125, 126, 129, 435, 440–442, 451 acetoclastic, 136 Methanogens (see also individual names), 57–59, 67–71, 73, 74, 181, 222, 223, 225, 227 Methanol, 58, 59, 82 as methyl donor, 69–71 catabolism, 67 -coenzyme M methyltransferase, 80, 81 Methanopyrus kandleri, 222, 223, 227–229 Methanosarcina sp., 59, 67–71, 73, 222 acetivorans, 67, 80 barkeri, 76, 80, 81 thermophila, 73, 74, 76, 385–387 Methanothermobacter marburgensis (see also Methanobacterium thermoautotrophicum), 117, 223, 227 thermoautotrophicus, 223 Methionine S-adenosyl-, see S-Adenosylmethionine biosynthesis, see Biosynthesis [methyl-13C]-, 225 seleno-, see Selenomethionine Methionine synthase, 2, 34–36, 56, 72, 75–82, 84, 86, 87, 90, 92, 93, 426–430, 432–435 cobalamin-dependent, 60–67

Met. Ions Life Sci. 2009, 6, 461–496

484 [Methionine synthase] conformational states, 65 electron transfer in, see Electron transfer EPR studies, see EPR human, see Human mammalian, see Mammalian mutants, 64–67 reductase, see Reductases structure, 64 wild-type, 65, 67 Methylamines, 58, 59 catabolism, 67, 69 -coenzyme M methyltransferase, 80–82 Methylation of B12 derivatives, 25, 26, 34, 35, 37 list of methyl donors, 35 Ni(I)F430, 125–128 Methyl bromide, 118, 122–124, 126 13 CH3, 123 deuterated, 123 Methylcobalamin (see also Cobalamins), 2–4, 16, 25, 32, 33, 36, 37, 54, 60–63, 65–67, 71, 78, 82–84, 87, 420, 425, 428, 433, 434, 451 activation, 427–430 as cofactor, 34 base-off, 15, 66, 76 base-off/His-on, 37 Co(II), 22, 23 Co(III), 22, 26, 145 crystal structure, see Crystal structures -dependent methyltransferases, see Transferases kinetics, 23, 24 molecular mechanics calculations, 422 NMR studies, see NMR photolysis, 30 reactivity, 24 structure, 11–15, 55, 63 Methylcobinamides, 23, 32, 76, 434 Co(II), 23 Co(III), 33, 75 Methyl-coenzyme M (see also Coenzyme M), 57–59, 119, 125, 127–129, 435, 439–442, 451 formation, 123 reduction, 125 structure, 117

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX Methyl-coenzyme M reductase (from) (see also F430), 57, 115–129, 420, 437 absorption spectroscopy, see Absorption spectroscopy active site, see Active site catalytic mechanism, 123, 124, 127–129, 419, 435–442, 451 conformational changes, 126 crystal structure, see Crystal structures ENDOR studies, see ENDOR spectroscopy EPR studies, see EPR hydrogen bonds, see Hydrogen bonds hyperfine sublevel correlation spectroscopy studies, see Hyperfine sublevel correlation spectroscopy inhibitors, 119, 120, 123 magnetic circular dichroism studies, see Magnetic circular dichroism Methanothermobacter marburgensis, 117, 223, 227 methyl transfer, see Methyl transfer Ni(I), 123, 124, 129, 236, 436, 437, 439–442 Ni(II), 129, 436, 437, 439–442 Ni(III), 129, 437, 439–442 nickel-alkyl species, 439–442 nickel-carbon bond, see Nickel-carbon bond Raman spectroscopy studies, see Raman spectroscopy Methyleneglutarate mutase, 89, 90 radical rearrangement, 89 reaction mechanism, 89 Methyl iodide, 118–120, 123 Methylmalonic aciduria, 93 Methylmalonyl-coenzyme A mutase active site, see Active sites conformational change, 91 crystal structure, see Crystal structures human, see Human mammalian, see Mammalian radical formation, 90, 92 reaction mechanism, 90, 92 wild-type, 91 Methylmercury, 32 Methylobacterium sp., 78 extorquens, 93, 94

SUBJECT INDEX Methyltetrahydrofolate in corrinoid iron-sulfur protein, 73, 74, 76, 136 methionine synthase, 35, 37, 60, 61, 65, 66, 70 2-(Methylthio)ethanesulfonate, see Methylcoenzyme M Methyl transfer (in), 75, 419 ACS/CODH, 138–140 catalysis, 71 cobalamins, 24, 25, 32, 62, 63, 432, 434, 435 cobamides, 38 coenzyme M, 68–71, 73 cofactors, 35 corrinoids, 31, 59, 139 corrins, 36 O-demethylase, 77 fosfomycin biosynthesis, 82–84 list of methyl donors, 69 methyl-coenzyme M reductase, 128, 441 Methyltransferases (in), 83 ACsE, 76 B12-dependent, 16, 35–38 coenzyme M, 58, 59, 67, 68, 78, 80, 81 conformational changes, 59, 82 corrinoid-dependent, 55–84, 136 O-demethylase, 77, 78 membrane-associated, 72 metabolism, see Metabolism Methanosarcina spp., 67–71 methylamine complex, 82 methylcobalamin-dependent, 432, 434 modes of action, 79–82 non-energy conserving cytoplasmic corrinoid, 72 reductive activation, 79–81 zinc in, 70, 71 Microbes anaerobic, 35 Microorganisms (see also individual names and species), 181 aerobic, 447 anaerobic, 447 anoxic environment, 375 growth, 365 nitrogen source, 366 Minerals iron-sulfur, 181, 199, 200 MoCu CODH, see Molybdenum-copper carbon monoxide dehydrogenase

485 Molecular mechanics calculations, 421, 422 Amber 95 force field, 422 F430, 437 quantum mechanics/-, see Quantum mechanics Molecular switch B12-binding nucleotides, 40 base-on/base-off in coenzyme B12, 5, 16–20 riboswitch, see Riboswitch Molluscs (see also individual names) hemocyanin, 299, 301, 302, 304–306, 316 Molybdenum (oxidation state undefined) 95 Mo, 397 97 Mo, 397 Molybdenum(IV), 332–334, 399, 400, 403 Molybdenum(V) (in) ‘‘rapid’’ species, 396, 397 ‘‘slow’’ species, 396, 397 synthetic model complexes for MoCu CODH, 334 ‘‘very rapid’’ species, 396, 397 xanthine oxidoreductase, 395–413 Molybdenum(VI), 332–334, 403 Molybdenum-copper carbon monoxide dehydrogenase (from), 298, 332–324, 343, 350 active site, see Active site density functional theory calculations, see Density functional theory calculations mechanism, 334 Oligotropha carboxidovorans, 332 structure, 332, 333 synthetic Mo(V) model, 334 X-ray absorption spectroscopy studies, see X-ray absorption spectroscopy Molybdenum enzymes (see also individual names), 395–413 Molybdenum nitrogenases, 370, 371, 373 P-cluster, 370 Molybdopterin (see also Pterin), 332, 333, 398 Monooxygenases cyanide in, 366 dopamine b-, see Dopamine b-monooxygenase methane, 378 peptidylglycine a-hydroxylating, see Peptidylglycine a-hydroxylating monooxygenase

Met. Ions Life Sci. 2009, 6, 461–496

486

SUBJECT INDEX

Moorella sp., 72 thermoacetica, 56, 73–77, 80, 134, 442 Mo¨ssbauer spectroscopy (studies of) ACS/CODH, 139–142, 146 corrinoid iron-sulfur protein, 56 [Fe]-hydrogenases, 229, 232, 233 [FeFe]-hydrogenases, 189, 192, 193, 450 iron guanylylpyridinol cofactor, 224, 225 nitrogenases, 371 mRNA B12 riboswitch, 18 Multi-copper oxidases active site, see Active sites blue, 338–340 cyanide in, 339, 340 intramolecular electron transfer, 339 Multiple anomalous dispersion, 228 Mutagenesis, 200 site-directed, see Site-directed mutagenesis Mutases, 85 amino-, see Aminomutases carbon skeleton, 38 glutamate, see Glutamate mutase isobutyryl-coenzyme A, see Isobutyrylcoenzyme A mutase MeaA, 94, 95 methyleneglutarate, see Methyleneglutarate mutase methylmalonyl-coenzyme A, see Methylmalonyl-coenzyme A mutase Myoglobin, 249, 251, 253, 257, 302, 326, 367 carbon monoxide complex, 244, 257 human, 301 isocyanide binding, 317 mammalian, 251

N NAD+, 171, 396 NADH oxidation, 184 NADPH, 79, 92, 104, 106, 245, 247, 257, 366, 430 Neisseria meningitidis, 248, 250, 255–258, 262, 274 Neurodegenerative diseases, 330 Neurological disoders, 387 Neurospora crassa, 304, 307 Neurotransmission, 245

Met. Ions Life Sci. 2009, 6, 461–496

Neutron Laue crystallography study of cob(II)alamin, 10 Nickel (oxidation state undefined) 61 Ni, 139, 444, 445 Nickel(0), 57 acetyl-coenzyme A synthase, 446, 447 ACS/CODH, 137, 138, 144–146 Nickel(I) (in), 57 acetyl-coenzyme A synthase, 58, 445, 447 ACS/CODH, 137, 139, 144–146 F430, see F430 F430M, see F430M methylation, 126 octaethylisobacteriochlorin, 120 redox couples, see Redox potentials Nickel(II) (in) ACS/CODH, 133–146, 444 alkyl species, 120, 123 F430, see F430 F430M, see F430M [NiFe]-hydrogenases, 169–171 redox couples, see Redox potentials –sulfur bond, see Bonds Nickel(III) (in), 117 acetyl-coenzyme A synthase, 446 ACS/CODH, 139 alkyl species, 120, 122, 123, 439–442 F430, see F430 F430M, see F430M hydride, 236 [NiFe]-hydrogenases, 168–172 redox couples, see Redox potentials thiolate, 436, 439, 440 Nickel-carbon bonds (in/with) acetyl groups, 134, 140, 141 ACS/CODH, 133–146 acyl-coenzyme A synthase, 59, 73, 76 carbon dioxide, 145 carbon monoxide, 144, 145 carbonyl groups, 134, 139, 140 carboxylate groups, 134, 143 dissociation energy, 120 F430, 125 methyl groups, 134–140, 144 methyl-coenzyme M reductase, 115–129 Nicotinamide, 28 adenine dinucleotide, see NAD+ adenine dinucleotide (reduced), see NADH adenine dinucleotide phosphate (reduced), see NADPH

SUBJECT INDEX [NiFe]-hydrogenases (from), 151–173, 180, 182, 184, 185, 190, 191, 201, 202, 220, 221, 234 active site, see Active sites Allochromatium vinosum, 158 carbon monoxide in, see Carbon monoxide carbon monoxide insertion, 161–164 catalytic mechanism, 153, 154, 222, 236 clusters, see Clusters conformational change, 156 crystal structure, see Crystal structures cyanide in, 158, 160–164, 171 density functional theory calculations, see Density functional theory calculations Desulfovibrio desulfuricans, 166 Desulfovibrio fructosovorans, 165–169 Desulfovibrio gigas, 153, 155–160, 166, 168, 187 Desulfovibrio vulgaris, 153, 156, 158, 168, 172 electron transfer in, see Electron transfer ENDOR studies, see ENDOR EPR studies, see EPR EXAFS studies, see Extended absorption fine structure spectroscopy F420-reducing, 222 [Fe2S4] cluster, 155, 156, 162, 165 Fourier transform infrared spectroscopy studies, see Fourier transform infrared spectroscopy hydrogen bonds, see Hydrogen bonds hyperfine sublevel correlation spectroscopy studies, see Hyperfine sublevel correlation spectroscopy studies of inactive states of the activ site, 168–171 infrared studies, see Infrared spectroscopy iron insertion, 161–164 maturation, 160–165 nickel insertion, 164 proton transfer, 166–168 redox potentials, see Redox potentials regulatory, 172 structure, 153–160 substrate binding, 172, 173 X-ray diffraction spectroscopy studies, see X-ray diffraction spectroscopy

487 [NiFeSe]-hydrogenases (from), 157, 165 crystal structure, see Crystal structures Desulfomicrobium baculatum, 156, 166 EXAFS studies, see Extended absorption fine structure spectroscopy Methanococcus voltae, 155, 156 Nitric oxide (in), 342, 380, 381 heme oxygenase complex, 255 inhibition of cytochrome c oxidase, 329, 330 oxygen evolving complex, 382, 383 reductase, see Reductases superoxide reductase, 381 synthase, see Synthases Nitrile, 365 Nitrite, 341, 342 Nitrite reductases, 340–342 active site, see Active sites blue, 341 carbon monoxide in, see Carbon monoxide copper in, 340–342 Cu(I), 341, 342 Cu(II), 341 green, 341 hydrogen bonds, see Hydrogen bonds inhibitors, 341 mutant, 341 structure, 341, 342 synthetic model complexes, 341, 342 wild-type, 341 Nitrogen (different oxidation states) (see also Dinitrogen) 14 N label, 331 15 N label, 56, 96, 158, 271, 272, 275, 331 15 N NMR, see NMR fixation, 200, 370 oxides, 298 Nitrogenases, 200, 201 EPR studies, see EPR magnetic circular dichroism studies, see Magnetic circular dichroism molybdenum, see Molybdenum nitrogenases Mo¨ssbauer spectroscopy studies, see Mo¨ssbauer spectroscopy sequences, 80 vanadium, see Vanadium nitrogenases X-ray absorption spectroscopy studies, see X-ray absorption spectroscopy

Met. Ions Life Sci. 2009, 6, 461–496

488 Nitrogen monoxide, see Nitric oxide Nitrous oxide reductase, 342–344 active site, see Active sites structure, 343 NMR (studies of) 1 H, 13, 14, 120, 262, 264, 265, 267, 274, 283, 284, 345, 348 2 H, 120 13 C, 13, 14, 259–264, 345, 367, 348, 372 15 N, 14, 266, 267, 271, 272 31 P, 14 B12 derivatives, 13 bleomycin, 349 [Fe]-hydrogenases, 229 glutamate synthase, 87 heme oxygenase, 249, 256, 260–267, 269, 271, 272, 274, 278–281, 283, 284 methylcobalamin, 11, 26 methyl-Ni(II)F430M, 120 relaxation measurements, 269–274 two-dimensional, 348, 349 NMRD, see Nuclear magnetic relaxation dispersion Non-heme iron enzymes, 377–388 2-His-1-carboxylate facial triad motif, 378–382 active site, see Active sites catalytic cycle, 379 cyanide interaction, 381–383 functions, 377, 378 structures, 377, 378 Norepinephrine production, 308 Nuclear magnetic relaxation dispersion measurements of galactose oxidase, 337 Nuclear magnetic resonance, see NMR Nuclear Overhauser effect spectroscopy studies of cob(III)yrinate, 14 methylcobalamin, 15 Nuclease P1, 386 Nucleophile or nucleophilic attack (by/in), 31, 32, 36, 37, 71, 237, 334 cob(I)alamin, 428 coenzyme M, 122 hydroxide, 137, 143, 385 nickel, 145, 439–441 substitution, 125–127 super-, 30

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX [Nucleophile or nucleophilic attack (by/in)] water, 369 xanthine oxidoreductase, 403, 409, 410 Nyctotherus ovalis, 184

O Octopus dofleini, 303 vulgaris, 302 Olefin Cu(I) complex, 298, 345 Olfactory receptor, 345, 346 Oligotropha carboxidovorans, 332 Operons cus, 344 hyp, 161, 162 nik, 161 Orconectes rusticus, 305 sanborni, 305 Origin of life, 181, 199 Ornithine 4,5-aminomutase, 95, 103 Oxidases (see also individual names) aldehyde, 368 blue multi-copper, 338–340 copper amine, see Copper amine oxidase cytochrome bo3, 327 cytochrome c, see Cytochrome c oxidase galactose, see Galactose oxidase heme-copper, see Heme-copper oxidase non-blue copper, 334–337 xanthine, see Xanthine oxidase Oxidative stress, 327, 330 Oxidoreductases, 383 aldehyde, 398, 403, 411 hydrogen:NAD(P)+, 165 quinoline-2-, 398 xanthine, see Xanthine oxidoreductase Oxygen (see also Dioxygen) 17 O EPR, see EPR 17 O label, 397, 399 18 O, 84, 300 evolving complex, see Oxygen evolving complex molecular, 153, 169, 244, 335 tolerance in bacteria, 171 Oxygenases heme-, see Heme oxygenasess mono-, see Monooxygenases

SUBJECT INDEX

489

Oxygen evolving complex (in) algae, 381 cyanide in, 382, 383 electron transfer in, see Electron transfer EPR studies, see EPR manganese, 373, 374 nitric oxide in, 382, 383 non-heme iron center, 380, 381 plants, 381 structure, 373 Oxyhemocyanin, 299

P Palladium in ACS/CODH, 146 Panulirus interruptus, 302 Paracoccus denitrificans, 318, 324 Penicillin, 378, 386 Peptidases (see also individual names) amino-, see Aminopeptidases endo- 386 exo-, 386 Peptidylglycine a-hydroxylating monooxygenase, 308–317 active site, see Active sites carbon monoxide in, see Carbon monoxide density functional theory calculations, see Density functional theory calculations electron transfer in, see Electron transfer EXAFS studies, see Extended absorption fine structure spectroscopy hydrogen bonds, see Hydrogen bonds infrared studies, see Infrared spectroscopy mutant, 313, 315, 316 structure, 309–311 truncated, 309–317 wild-type, 313 X-ray absorption spectroscopy studies, see X-ray absorption spectroscopy Peptidyl prolyl cis/trans isomerase SlyD, 164 Perchloroethylene, 77, 78 Peroxidases, 252, 253. 374 bromo-, see Bromoperoxidase chloro-, see Chloroperoxidase halo-, see Haloperoxidase iodo-, see Iodoperoxidase microperoxidase-8, 170

Peroxide(s), 168, 252 hydro-, see Hydroperoxides hydrogen, see Hydrogen peroxide Peroxynitrite, 330 1,10-Phenanthroline, 140, 445 Phenolase activity, 299, 304 tyrosinase, 303 Phenolates (or phenols and phenolic groups), 299, 339 hydroxylation, 304 ortho-di-, 299 polychlorinated, 78 Phosphate, 368 inorganic, 59, 369 meta-, see Metaphosphate pyridoxal, 102, 103 seleno-, 162 Phosphatases acyl-, 162 alkaline, see Alkaline phosphatase hydrogen bonds, see Hydrogen bonds lipid, 369 Phospholipase C, 386 Phosphonate hydroxyethyl-, 83, 84 Phosphonoacetaldehyde, 83 Phosphorescence, 306 Photolysis (of), 73 laser flash, 300, 339 methylcobalamin, see Methylcobalamin Photosynthesis, 182, 337 Photosystem II, 380, 381 oxygen evolving complex, see Oxygen evolving complex water oxidizing complex, 373 Photothermal beam deflection, 320 Phycobilins, 246 Phylogenetic analysis, 184 Phytochromes, 283, 284 bacterio-, 283, 284 Pisum sativum, 376 Plants carbonic anhydrase, 385 heme oxygenase in, see Heme oxygenase nitrogen source, 366 oxygen evolving complex, 381 photosynthetic, 373 Plastocyanin, 338 Platinum in ACS/CODH, 146

Met. Ions Life Sci. 2009, 6, 461–496

490 Porphyrins (see also Hemes and individual names), 267 Fe(III), 252, 259, 261, 367 hydro-, 116 nickel, 116 radical, see Radicals tetraphenyl-, 259 Positron emission tomography, 350 Potassium (in), 70, 71 diol dehydrase, 96, 98–100 Prebiotic chemistry, 181, 199, 206 Prokaryotes (see also individual names), 54, 56, 60, 84, 334, 374 carbonic anhydrase, 384, 385 non-photosynthetic, 284 1,2-Propanediol, 95–98 [1-3H]1,2-, 84 18 O-enriched, 84 dehydrase, see Dehydrases Propionaldehyde, 84 Propionibacterium shermanii, 90, 101 Proteases (see also individual names) nickel-dependent, 161 Proteins (see also Enzymes and individual names) copper, see Copper proteins DNA-binding, 161 Hyc, 162 Hyd, 201, 203, 204 Hyp, 162–164 methyltransferase-activating, 80–82 molybdenum iron, 370, 371 NikR, 161, 223 Ram, 80, 82 structural probing, 295–350 vanadium iron, 370, 371 Protein Data Bank (codes of protein structures) ascorbate oxidase, 340 heme oxygenases, 249, 270, 272, 278, 282 [NiFe]-hydrogenases, 166 Protists, 181 Protocatechuate 3,4-dioxygenases, 380, 382 active site, see Active sites crystal structure, see Crystal structures cyanide in, 380, 382 hydrogen bonds, see Hydrogen bonds Proton transfer (in), 39 cytochrome c oxidases, 323, 324 heme-copper oxidases, 326, 327 [NiFe]-hydrogenases, 166–168

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX Prussian blue, 365, 388 Pseudomonas aeruginosa, 248, 250, 252, 256–258, 260–284, 338 Pteridines 2-amino-4-oxo-, see Pterin 2,4,7-trihydroxy-, see Violapterin 2,6-dihydroxy-, see Lumazine Pterins methenyltetrahydromethano-, 221, 222, 229, 235–237 methylenetetrahydromethano-, 220–222, 236 methyltetrahydro-, 35, 59 methyltetrahydromethano-, 58, 73, 80, 152 methyltetrahydrosarcino-, 73 molybdo-, see Molybdopterin pyrano-, see Pyranopterin viola-, 408 Purines (see also individual names) 2,6-diamino-, 410 2-hydro-6-methyl-, 399, 401–403, 406–410 6-mercapto-, 404 metabolism, see Metabolism Pyranopterin cofactor molybdenum enzymes, 398 structure, 398 tungsten enzymes, 398 Pyrimidine 4,6-dihydroxypyrazolo[3,4-d]-pyrimidine, see Alloxanthine 4-hydroxypyrazolo[3,4-d]-pyrimidine, see Allopurinol Pyruvate carboxylase, 383 Pyruvate formate lyase activating enzyme, 203, 205

Q Quantum mechanics/molecular mechanics calculations, 166, 167, 275, 422–424, 432, 433, 451 Quercetinase, 298 Quinones, 380 ortho-, 299

R Radicals (see also individual names) acetyl, 30 adenosyl, 24, 84, 88, 101, 431, 432

SUBJECT INDEX [Radicals (see also individual names)] S-adenosylmethionine, 201–207 alkyl, 29, 31, 40, 349 allyl, 128 amino acid, 88, 89, 203, 205, 206, 337, 378 cyclopropinyl, 89 cyclopropylcarbinyl, 90 cysteine, 104, 105 5 0 -deoxy-5 0 -adenosyl, 29, 38–40, 91, 92, 96, 97, 104–106 disulfide, 124, 125, 127 ethyl, 128 ferrous keto p neutral, 257 glutamyl, 88 glycine, 206 glycyl, 88, 89 hydroxyl, 267, 275 methyl, 22, 23, 33, 36, 124, 128, 439–442 b-methylaspartate, 88 porphyrin, 267 reactions, 30, 31 rearrangement, 88, 89, 419 redox couples, see Redox potentials S–CoB, 439–441 semiquinone, 335 succinyl-coenzyme A, 92 thiyl, 104–106, 124, 125, 128, 436 ‘‘trap’’, 29, 30 trichlorovinyl, 35, 37 tyrosyl, 337, 378 Radiolabels, 350 Ralstonia cuprivorans, 171 eutropha, 171, 172 Raman spectroscopy (studies of) cobalamins, 425 copper-zinc superoxide dismutase, 368 corrinoid iron-sulfur protein, 75 heme-copper oxidases, 324, 332 methyl-coenzyme M reductase, 437 UV, 332, 345 Rat heme oxygenase, see Heme oxygenases Rate constants association, 251, 258, 300, 301 B12 derivatives, 23 carbon monoxide transfer, 322, 323 cobalamins, 91, 92 dissociation, 251, 258, 301, 302, 328, 339 isotope exchange, 269–271 methyl transfer, 139, 140

491 Reactive oxygen species (see also individual names), 234, 327 detoxification, see Detoxification toxicity, see Toxicity Redox potentials (in) ACS/CODH, 144 B12 derivatives, 19, 20, 26 blue copper proteins, 338 Co(II)/Co(I), 20–23, 26, 29, 78, 429, 430 Co(III)/Co(II), 20–23 cob(II)alamin, 429, 430 cob(II)alamin/cob(I)alamin, 61, 62 cob(II)amide/cob(I)amide, 75 cob(II)inamide, 429, 430 coenzyme F430, 118 Cu(II)/Cu(I), 305, 337 disulfide/disulfide radical, 124 [Fe4S4]2+/[Fe4S4]–, 172 Fe(III)/Fe(II), 325, 326 halide oxidation, 372 heme-copper oxidases, 325, 326 laccase, 339 methyl-Ni(III)F430M/methyl-Ni(II)F430M, 120 NADPH, 429 Ni(II)F430M/Ni(I)F430M, 117, 120 Ni(III)F430M/Ni(II)F430M, 117 [NiFe]-hydrogenases, 159, 165, 169 quinone/semiquinone, 61 semiquinone/hydroquinone, 61 thyil radical/thiol, 124 Redox processes B12 derivatives, 18–24 kinetics, 22–24 thermodynamics, 20–22 Reductases alkyne, 372 coenzyme M, 58 cytochrome P450, 247, 257 ferredoxin, 79, 81, 248 flavodoxin, 79, 81 isocyanide, 372 methionine synthase, 81, 92, 93 methyl-coenzyme M, see Methylcoenzyme M reductases nitric oxide, 330 nitrite, see Nitrite reductase nitrous oxide, see Nitrous oxide reductase ribonucleotide triphosphate, see Ribonucleotide triphosphate reductase

Met. Ions Life Sci. 2009, 6, 461–496

492

SUBJECT INDEX

[Reductases] ribonucleotide, 38 superoxide, see Superoxide reductase thioredoxin, 104, 106 Reduction potentials, see Redox potentials Resonance Raman spectroscopy, see Raman spectroscopy Respiration, 182, 366 inhibition, 327 Rhodanese, 366 Rhodobacter capsulatus, 398, 403–408, 410 sphaeroides, 318, 320, 321, 324 Rhodospirillum rubrum, 136, 141 Ribonucleic acid, see RNA Ribonucleotide reduction, 124 Ribonucleotide triphosphate reductase, 103–106, 378 crystal structure, see Crystal structures electron transfer in, see Electron transfer reaction cycle, 104, 105 Riboswitch in B12, 18, 40 RNA, 16 hydrolysis, 386 m-, see mRNA polymerase, 383

S Salmonella typhimurium, 101, 161 Seaweed bromoperoxidase, 369 Selenium [NiFeSe]-hydrogenases, see [NiFeSe]hydrogenases Selenocysteine in [NiFeSe]-hydrogenases, 156, 166 Selenomethionine, 228 Selenophosphate synthase, 162 Self-consistent field method complete active space, 423, 426 coupled-perturbed, 425, 445 multiconfigurational, 422, 423, 451 zeroth order regular approximation, 445 Semiquinones, 383 lumiflavin, 339 Siderophores, 161

Met. Ions Life Sci. 2009, 6, 461–496

Site-directed mutagenesis of blue-copper proteins, 338 [Fe]-hydrogenases, 231 heme oxygenase, 249, 279 [NiFe]-hydrogenases, 156 peptidylglycine a-hydroxylating monooxygenase, 316 SlyD, 164 Smell receptors, 345, 346, 350 Snake venom, 388 Sodium, 72 borohydride, 122 dithionite, 188, 383 transport, 106 Sporomusa ovata, 56, 59 Stability constants (see also Association constants, Equilibrium constants, and Inhibition constants) carbon monoxide binding in heme-copper oxidases, 330 carbon monoxide binding in hemocyanins, 302 carbon monoxide binding in synthetic Cu(I) carbonyl complexes, 300, 301 dioxygen binding in hemocyanins, 302 dioxygen binding in synthetic Cu(I) carbonyl complexes, 301 hydrogen peroxide binding in manganese catalases, 376 Stacking p-, 306, 307 tryptophan/tyrosine, 337 tyrosine/histidine, 313 Steady-state kinetics ACS/CODH, 140 methyl-coenzyme M reductase, 439 Stopped flow kinetics ACS/CODH, 139 Streptomyces sp., 83, 303 cinnamonensis, 94 collinus, 94 glaucescens, 307 verticillus, 347, 348 Sulfate, 126, 165 Sulfenate in [NiFe]-hydrogenases, 169–171 Sulfide (in), 118 (m)-bridging, 134–136, 142, 145, 332, 334, 343 ACS/CODH, 143

SUBJECT INDEX [Sulfide (in)] di-, see Disulfide [NiFe]-hydrogenases, 159 Sulfonamides, 387 Sulfonic acid (or sulfonate) 2-bromoethane, 123, 126 3-bromopropane-, see 3Bromopropanesulfonate ester hydrolysis, 385 propane, 122 Sulfur 33 S, 397 –Ni(II) bond, see Bonds Sulfurospirillum multivorans, 34 Superoxide, 330, 381 Cu(II)-, see Copper(II) dismutase, see Superoxide dismutase disproportionation, 374 [Fe]-hydrogenase inactivation, 234, 235 reductase, see Superoxide reductase reduction, 381 scavenging, 331 Superoxide dismutase(s), 330, 373, 381 Cu-Zn, see Copper-zinc superoxide dismutase Escherichia coli, 376 iron, 376 manganese, see Manganese superoxide dismutase zinc, 383 Superoxide reductases, 379, 381 active site, see Active sites azide in, see Azide cyanide in, 381, 382 electron density, see Electron density electron transfer in, see Electron transfer EPR studies, see EPR magnetic circular dichroism studies, see Magnetic circular dichroism nitric oxide in, see Nitric oxide Surface plasmon resonance studies of heme oxygenase, 247 Synchrotron radiation, 157 Synechocystis PCC6803, 246 Synthases acetyl-coenzyme A, see Acetyl-coenzyme A synthase aminoacyl-tRNA, 383 aminoimidazole ribonucleotide, 162 biotin, see Biotin synthase

493 [Synthases] carbamoyl phosphate, see Carbamoyl phosphate synthase deacetoxycephalosporin C, 378 glutamate, 87 isopenicillin N, 378 lipoate, 203 methionine, see Methionine synthase nitric oxide, 244 selenophosphate, 162 thiazole, 203

T Tartaric acid, 87 Testis, 245 Tetrachloroethylene, 35, 78 Tetrahydrofolate, 37, 57, 60, 72, 77, 78, 80 methyl-, see Methyltetrahydrofolate Tetrahydromethanopterin, 221 methenyl-, 221, 222, 229, 235–237 methyl-, 58, 73, 80, 152 methylene-, 220–222, 236 Tetrahydropterins, 57, 58 Tetramethylammonium ion, 69 Tetrapyrroles, 117, 246 F430, see F430 Thallium(I) in diol dehydrase, 98 Thermal springs, 199 Thermolysin, 383 Thermotoga maritima, 63, 64, 186 Thermus thermophilus, 318, 324, 327, 375, 376 Thiocarbamates, 333, 365 Thiocyanate (in), 365, 368, 372, 387, 397 ACS/CODH, 135, 142 oxidation, 372 Thioether bridge, 303, 306 N-7-Thioheptanoyl-O-phospho-L-threonine, see Coenzyme B Thiols (and thiolate groups) (in) (see also Mercapto-), 35, 104, 122, 125, 126 ACS/CODH, 135, 136, 145 alkyl transfer, 71 bridging, 145 catabolism, 67 di-, see Dithiol [FeFe]-hydrogenases, 182, 183, 185, 187 iron guanylylpyridinol cofactor, 226

Met. Ions Life Sci. 2009, 6, 461–496

494 [Thiols (and thiolate groups) (in) (see also Mercapto-)] methyl-, 58, 59, 67, 69 2-(methylthio)ethanesulfonate, see Methyl-coenzyme M Ni(II), 436 [NiFe]-hydrogenases, 155, 157, 162, 163, 166 Thioredoxins, 104, 106 Titanium(III) reduction of Ni(II)F430, 120, 122 Titanium(III) citrate in ACS/CODH, 137 methyl-coenzyme M reductase, 118 Toxicity of carbon monoxide, 243–245 cyanide, 330, 365 Transfer alkyl, 71 electron, see Electron transfer hydride, see Hydride transfer hydrogen, 101 hydroxyl group, 403 methyl, see Methyl transfer proton, see Proton transfer Transferases (see also individual names) adenosyl-, 34, 92, 93 ATP:corrinoid adenosyl-, 426, 427, 429, 451 methyl-, see Methyltransferases O-carbamoyl, 162 zinc, 383 Transport(ers) (of) ABC-type, 161 B12, 18 copper, 340 iron, 161 nickel, 161, 164 Triclosan oxidation, 340 2,4,5-Trihydroxylphenylalanine quinone, 334–337 Trimethylphosphine, 198 Tris, see Tris(hydroxymethyl)methylamine Tris(hydroxymethyl)methylamine buffer, 223, 234 Tris(pyrid-2-ylmethyl)amine Cu(I) comples, 300–302 Tritium, 84, 96 label, 104 Trypsin, 369

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX Tryptophan, 337, 344, 345 Tungsten enzymes, 398 Tyrosinases, 298, 299, 304–308 active site, see Active sites bacterial, 307 fungal, 303, 304, 307 mammalian, 304 reaction cycles, 303 Tyrosine, 313, 337 radical, 337, 378

U Urea, 385 Urease, 164 Uric acid, 400, 404 Uridine triphosphate, see UTP UTP, 104 UV absorption spectrocopy (studies of) (see also Absorption spectroscopy), 28, 226, 332, 345 ACS/CODH, 139 B12 derivatives, 20 iron guanylylpyridinol cofactor, 226 Vis, 139, 226

V Vanadium (different oxidation states) (in) biology, 368 chloroperoxidase, see Chloroperoxidases nitrogenases, see Vanadium nitrogenases Vanadium(V), 368–370 Vanadium enzymes (see also individual names), 368–373 Vanadium nitrogenases, 368–373 Azotobacter, 370 carbon monoxide interaction, 373 hydrazine release, 372 Vanillate O-demethylase, 77 Vasodilation carbon monoxide in, 245 Vertebrates, 299 Violapterin, 408 Vitamin B12 (and derivatives) (see also Coenzyme B12), 1–41, 365 atom numbering, 7 deficiency, 426 electron transfer, see Electron transfer

SUBJECT INDEX

495

[Vitamin B12 (and derivatives) (see also Coenzyme B12)] history, 2 neo-, 7, 8, 26 nor-, 8 norpseudo-, 35 pseudo-, 4, 8, 9, 16, 26, 35, 54, 55 structure, 3–18 Voltammetry cyclic, see Cyclic voltammetry

W Walterinnesia aegyptia, 388 Wastewater, 340, 366 Water 17 O-labeled, 397, 399 oxidation, 373 Wood-Ljungdahl pathway, 56–60, 72, 73, 76

X Xanthine 8-[13C]-, 397, 400 8-[2H]-, 397 allo-, see Alloxanthine as substrate, 400–402 dehydrogenase, see Dehydrogenases hydroxylation, 412 tautomeric forms, 412, 413 Xanthine oxidase, 234, 368 desulfo, 413 FYX-051 complex, 405, 406 Xanthine oxidoreductase (from), 395–413 active site, see Active sites alloxantine-complexes, 404, 405 bovine, 398, 402, 403, 405–409, 411 complexed with FYX-051, 405, 406 crystal structure, see Crystal structures cyanide in, 397 desulfo, 406, 407, 409, 410 electron transfer in, see Electron transfer ENDOR studies of the ‘‘very rapid’’ species, 400–405 EPR studies, see EPR ESEEM studies, see Electron spin echo envelope modulation hydrogen bonds, see Hydrogen bonds

[Xanthine oxidoreductase (from)] 2-hydroxy-6-methylpurine as substrate, 401–403 2-hydroxy-6-methylpurine complex, 406–409 inhibition, 404–406 lumazine complex, 406–409 mammalian, 405 model complexes, 403 molybdenum center, 411 mutant, 410–412 reaction mechanism, 395–413 Rhodobacter capsulatus, 398, 403, 405–408, 410 substrate orientation, 410–413 wild-type, 410–412 xanthine as substrate, 400, 401 X-ray absorption spectroscopy studies, see X-ray absorption spectroscopy XAS, see X-ray absorption spectroscopy Xenobiotics, 28 X-ray absorption spectroscopy (studies of) copper amine oxidase, 334, 337 dopamine b-monooxygenase, 311, 314, 315 [Fe]-hydrogenases, 229, 232, 233 iron guanylylpyridinol cofactor, 224, 225 MoCu CODH, 332 nitrogenases, 371 peptidylglycine a-hydroxylating monooxygenase, 311, 312, 317 xanthine oxidoreductase, 397–399, 411 X-ray crystal structure studies (of) (see also Crystal structures) ACS/CODH, 443, 444 bleomycin, 347, 348 catechol oxidase, 303 copper amine oxidase, 335 copper-zinc superoxide dismutase, 331 cytochrome bo3 oxidase, 327 cytochrome c oxidase, 323 deoxyhemocyanin, 299 diol dehydrase, 96 [FeFe]-hydrogenases, 183, 189, 192, 447, 448 [Fe]-hydrogenase, 228–230 heme-copper oxidases, 318, 321 heme oxygenase, 248, 249, 253, 255, 256, 270, 276, 279, 282 manganese catalase, 375

Met. Ions Life Sci. 2009, 6, 461–496

496 [X-ray crystal structure studies (of) (see also Crystal structures)] methylmalonyl-coenzyme A mutase, 90, 92 MoCu CODH, 332 [NiFe]-hydrogenases, 153–158, 162, 172 nitrite reductase, 341, 342 nitrogenases, 370, 371 norpseudovitamin B12, 9 oxygen evolving complex, 373 oxyhemocyanin, 299 peptidylglycine a-hydroxylating monooxygenase, 309–311 superoxocob(III)alamin, 10 synthetic Cu(II)-carbonyl models, 316, 317, 322 tyrosinase, 303 vanadium chloroperoxidase, 369 vitamin B12, 7 X-ray diffraction spectrocopy (studies of) carbon monoxide dehydrogenase, 141–143, 146 [FeFe]-hydrogenases, 190, 191 [Fe]-hydrogenases, 232 heme oxygenase, 249

Met. Ions Life Sci. 2009, 6, 461–496

SUBJECT INDEX [X-ray diffraction spectrocopy (studies of)] iron guanylylpyridinol cofactor, 225 [NiFe]-hydrogenases, 157

Z Zinc (different oxidation states) (in) biology, 383–386 catalytic, 70, 71 Zinc(II) (in) ACS/CODH, 142 alkaline phosphatase, 388 Zinc enzymes (see also individual names), 383–388 active sites, see Active sites cyanide interaction, 387, 388 dinuclear, 386 structure, 384, 386 Zinc hydrolases (see also individual names), 383, 388 cyanide interaction, 388 Zucchini squash ascorbate oxidase, 339

Met. Ions Life Sci. 2009, 6, 497–510

Author Index of Contributors to MIBS and MILS Prepared to celebrate publication of the 50th volume edited by the Sigels: Metal Ions in Biological Systems; MIBS-1 to MIBS-44 (1973–2005) Metal Ions in Life Sciences; MILS-1 to MILS-6 (2006–2009)

A Abadin, H. G., MILS-1 (2006) 395–425 Abt, D. J., MIBS-39 (2002) 369–403 Achterberg, E., MILS-5 (2009) 441–481 Adams, M. W. W., MIBS-39 (2002) 673–697 Agarwal, R. P., MIBS-2 (1973) 167–206 Aime, S., MIBS-40 (2003) 643–682 Aisen, P., MIBS-35 (1998) 585–631 Akermann, B., MIBS-33 (1996) 177–252 Albrecht-Gary, A.-M., MIBS-35 (1998) 239–327 Alema`, S., MIBS-17 (1984) 275–317 Alessio, E., MIBS-42 (2004) 323–351 Allen, G., MIBS-38 (2001) 197–212 Allen, M. J., MIBS-42 (2004) 1–38 Altura, B. M., MIBS-26 (1990) 359–416

Altura, B. T., MIBS-26 (1990) 359–416 Alvarez, R., MIBS-16 (1983) 103–122 Ando, K., MIBS-19 (1985) 207–227 Andreesen, J. R., MIBS-39 (2002) 405–430 Andrews, R. K., MIBS-23 (1988) 165–284 Andrews, S. C., MIBS-35 (1998) 435–477 Andronikashvili, E. L., MIBS-10 (1980) 167–206; MIBS-23 (1988) 331–357 Anni, H., MIBS-28 (1992) 219–241 Antholine, W. E., MIBS-33 (1996) 619–648; MIBS-42 (2004) 463–497 Antonietti, M., MILS-4 (2008) 607–643 Antonini, E., MIBS-13 (1981) 187–228 Anzellotti, A., MIBS-41 (2004) 379–419 Aoki, K., MIBS-32 (1996) 91–134 Arai, F., MIBS-29 (1993) 137–160 Arakawa, Y., MIBS-29 (1993) 101–136 Arceneaux, J. E. L., MIBS-35 (1998) 37–66


498

AUTHOR INDEX

Arimura, K., MIBS-34 (1997) 405–420 Arpalahti, J., MIBS-32 (1996) 379–395 Artymiuk, P. J., MIBS-35 (1998) 435–477 Ash, D. E., MIBS-37 (2000) 407–428 Atkins, W. M., MIBS-25 (1989) 417–475 Atrian, S., MILS-5 (2009) 155–181 Atwood, C. S., MIBS-36 (1999) 309–364 Auld, D. S., MIBS-25 (1989) 359–394 Auling, G., MIBS-30 (1994) 131–161 Aull, J. L., MIBS-11 (1980) 337–376 Averill, B. A., MIBS-7 (1978) 127–183

B Babcock, G. T., MIBS-30 (1994) 77–107; MIBS-37 (2000) 613–656 Baccini, P., MIBS-18 (1984) 1–4; MIBS-18 (1984) 239–286; MIBS-18 (1984) 353–361 Baichoo, N., MIBS-38 (2001) 255–287 Balamurugan, K., MILS-5 (2009) 31–49 Baldi, F., MIBS-34 (1997) 213–257 Baldwin, G. S., MIBS-37 (2000) 345–364 Bansil, R., MIBS-7 (1978) 311–349 Bara, M., MIBS-26 (1990) 549–578 Baran, E. J., MIBS-31 (1995) 129–146; MILS-4 (2008) 219–254 Barge, A., MIBS-40 (2003) 643–682 Barnes, K. R., MIBS-42 (2004) 143–177 Barnett, M. O., MIBS-34 (1997) 185–212 Barton, J. K., MIBS-25 (1989) 31–103; MIBS-33 (1996) 325–365; MIBS-36 (1999) 211–249 Bartzokis, G., MILS-1 (2006) 151–177 Basile, L. A., MIBS-25 (1989) 31–103 Battell, M., MIBS-31 (1995) 575–594 Battigello, J.-M., MIBS-33 (1996) 593–617 Bau, R., MIBS-8 (1979) 1–55 Bayer, E., MIBS-31 (1995) 407–421 Bayer, T. A., MILS-1 (2006) 115–123 Bazylinski, D. A., MILS-4 (2008) 343–376 Beaulieu, R., MILS-2 (2007) 473–500 Beck, M. T., MIBS-7 (1978) 1–28 Beckie, R. D., MIBS-44 (2005) 145–169 Behra, R., MIBS-44 (2005) 47–73 Bell, S. G., MILS-3 (2007) 437–476 Beratan, D. N., MIBS-27 (1991) 97–127 Bereswill, S., MILS-2 (2007) 545–579 Berg, J. M., MIBS-25 (1989) 235–254 Berg, T., MIBS-44 (2005) 1–19

Met. Ions Life Sci. 2009, 6, 497–510

Bergamo, A., MIBS-42 (2004) 323–351 Berman, A., MILS-4 (2008) 167–205 Bernadou, J., MIBS-33 (1996) 399–426 Bernauer, K., MIBS-1 (1974) 117–128; MIBS-27 (1991) 265–289 Berndt, C., MILS-5 (2009) 413–439 Bernhardt, R., MILS-3 (2007) 361–396 Bertini, I., MIBS-12 (1981) 31–74; MIBS-15 (1983) 101–156; MIBS-21 (1987) 47–86 Beuerle, J., MIBS-28 (1992) 455–505 Beveridge, T. J., MILS-4 (2008) 127–165 Biggs, W. R., MIBS-6 (1976) 141–196 Billadeau, M. A., MIBS-33 (1996) 269–296 Bingham, F. T., MIBS-20 (1986) 119–156 Birch, N. J., MIBS-14 (1982) 257–313; MIBS-26 (1990) 105–117; MIBS-41 (2004) 305–332 Birkedal, H., MILS-4 (2008) 295–325 Birnbaum, E. R., MIBS-6 (1976) 251–290 Bishop, K. H., MIBS-34 (1997) 113–130 Blakeley, R. L., MIBS-23 (1988) 165–284 Blindauer, C. A., MILS-5 (2009) 51–81 Bloemink, M. J., MIBS-32 (1996) 641–685 Bodaly, R. A., MIBS-34 (1997) 259–287 Boddin, M., MIBS-31 (1995) 45–88 Bogumil, R., MIBS-37 (2000) 365–405 Bollag, J.-M., MIBS-28 (1992) 205–217 Bond, A. M., MIBS-27 (1991) 431–494; MILS-3 (2007) 127–155 Booth, B. P., MIBS-36 (1999) 723–749 Boskey, A. L., MILS-4 (2008) 457–505 Bosselmann, F., MILS-4 (2008) 207–217 Botta, M., MIBS-40 (2003) 643–682 Boudou, A., MIBS-34 (1997) 289–319 Boudvillain, M., MIBS-33 (1996) 87–104 Bourdon, R., MIBS-16 (1983) 245–260 Boyle, R. W., MIBS-23 (1988) 1–29 Bradley, P. M., MIBS-40 (2003) 323–353 Branchaud, B. P., MIBS-36 (1999) 79–102 Braun, V., MIBS-35 (1998) 67–145 Brautbar, N., MIBS-26 (1990) 285–320 Braze˙ nas, G., MIBS-10 (1980) 253–279 Bregadze, V. G., MIBS-23 (1988) 331–357; MIBS-32 (1996) 419–451; MIBS-33 (1996) 253–267 Breslow, E., MIBS-3 (1974) 133–155 Brewer, G. J., MIBS-14 (1982) 57–75 Briat, J.-F., MIBS-35 (1998) 563–584 Briggs, F. N., MIBS-6 (1976) 323–398 Bringmark, L., MIBS-34 (1997) 161–184 Brodin, P., MIBS-25 (1989) 255–307

AUTHOR INDEX

499

Brondino, C. D., MIBS-39 (2002) 539–570 Brown III, R. D., MIBS-21 (1987) 229–270 Brown, D. A., MIBS-14 (1982) 125–177 Brown, D. R., MILS-1 (2006) 89–114 Brown, K. C., MIBS-38 (2001) 351–384 Brown, N. L., MIBS-30 (1994) 405–434; MIBS-34 (1997) 527–568 Brown, R. S., MIBS-15 (1983) 55–99 Brunold, T. C., MILS-6 (2009) 417–460 Brunori, M., MIBS-13 (1981) 187–228 Bryngelson, P. A., MILS-2 (2007) 417–443 Buckingham, D. A., MIBS-38 (2001) 43–102 Buffle, J., MIBS-18 (1984) 165–221 Bugg, C. E., MIBS-17 (1984) 51–97 Bulman, R. A., MIBS-40 (2003) 39–67; MIBS-40 (2003) 683–706 Bult, A., MIBS-16 (1983) 261–278 Bu¨nzli, J.-C. G., MIBS-42 (2004) 39–75 Burger, D., MIBS-17 (1984) 215–273 Burger, K., MIBS-9 (1979) 213–250 Burgmayer, S. J. N., MIBS-39 (2002) 265–316 Burrows, C. J., MIBS-33 (1996) 537–560; MIBS-38 (2001) 289–311 Burstyn, J. N., MIBS-38 (2001) 103–143 Bush, A. I., MIBS-36 (1999) 309–364; MILS-1 (2006) 1–7; MILS-1 (2006) 427–435 Butler, A., MIBS-44 (2005) 21–46 Butler-Ransohoff, J. E., MIBS-25 (1989) 395–415 Butt, T. R., MIBS-25 (1989) 147–169 Buxton, G. V., MIBS-36 (1999) 103–123 Byers, B. R., MIBS-35 (1998) 37–66

C Campbell, P. G. C., MILS-5 (2009) 239–277 Carafoli, E., MIBS-17 (1984) 129–186 Carter, B. J., MIBS-33 (1996) 593–617 Caruso, F., MIBS-42 (2004) 353–384 Carver, P. L., MILS-3 (2007) 591–617 Casey, C. E., MIBS-16 (1983) 1–26 Chan, A. W. K., MIBS-37 (2000) 123–156 Chang, C. J., MILS-1 (2006) 321–370 Charleston, J. S., MIBS-34 (1997) 371–403 Chasteen, N. D., MIBS-31 (1995) 231–247; MIBS-35 (1998) 479–514 Chen, B., MIBS-44 (2005) 171–203 Cheng, Y., MIBS-40 (2003) 707–751

Chidambaram, M. V., MIBS-14 (1982) 125–177 Chiswell, B., MIBS-35 (1998) 667–690 Chkhaberidze, J. G., MIBS-33 (1996) 253–267 Chlebowski, J. F., MIBS-6 (1976) 1–140 Christensen, H. E. M., MIBS-27 (1991) 57–96 Christensen, J. M., MIBS-16 (1983) 185–199 Christianson, D. W., MIBS-37 (2000) 407–428 Chung, R., MILS-5 (2009) 279–317 Chutkow, J. G., MIBS-26 (1990) 417–440; MIBS-26 (1990) 441–461 Cillard, J., MIBS-36 (1999) 251–287 Ciurli, S., MILS-2 (2007) 241–277 Clark, C. R., MIBS-38 (2001) 43–102 Clarke, M. J., MIBS-11 (1980) 231–283; MIBS-32 (1996) 727–780 Classen, H.-G., MIBS-26 (1990) 321–339; MIBS-26 (1990) 597–609; MIBS-41 (2004) 41–69 Clayden, N. J., MIBS-21 (1987) 187–227 Cleare, M. J., MIBS-11 (1980) 1–62 Clegg, M. S., MIBS-37 (2000) 89–121 Cohen, L., MIBS-26 (1990) 271–284; MIBS26 (1990) 505–512 Colburn, R. W., MIBS-6 (1976) 291–321 Cole, M. M., MIBS-23 (1988) 47–90 Coleman, J. E., MIBS-6 (1976) 1–140; MIBS-25 (1989) 171–234 Co¨lfen, H., MILS-4 (2008) 607–643 Collison, D., MIBS-31 (1995) 617–670 Coluccia, M., MIBS-42 (2004) 209–250 Comte, M., MIBS-17 (1984) 215–273 Conrad, D. W., MIBS-27 (1991) 199–222 Conrad, L. S., MIBS-27 (1991) 57–96 Contakes, S. M., MILS-3 (2007) 157–185 Cooperman, B. S., MIBS-5 (1976) 79–126 Cosper, M. M., MIBS-39 (2002) 621–654 Costa, M., MIBS-20 (1986) 259–278; MIBS20 (1986) 279–303 Cox, J. A., MIBS-17 (1984) 215–273 Cox, J. D., MIBS-37 (2000) 407–428 Craig, P. J., MIBS-29 (1993) 37–77 Crans, D. C., MIBS-31 (1995) 147–209; MIBS-31 (1995) 287–324 Creutz, C. E., MIBS-17 (1984) 319–351 Crichton, R. R., MIBS-35 (1998) 633–665; MIBS-41 (2004) 185–219; MILS-1 (2006) 227–279

Met. Ions Life Sci. 2009, 6, 497–510

500

AUTHOR INDEX

Crooke, S. T., MIBS-25 (1989) 147–169 Crowley, J. D., MIBS-37 (2000) 209–278 Crumbliss, A. L., MIBS-35 (1998) 239–327 Cryle, M. J., MILS-3 (2007) 397–435 Cui, M., MIBS-33 (1996) 593–617 Culotta, V. C., MIBS-37 (2000) 35–56 Cunha, C. A., MIBS-39 (2002) 539–570 Curtis, N. J., MIBS-15 (1983) 55–99

D Dabrowiak, J. C., MIBS-11 (1980) 305–336 Dalbie`s, R., MIBS-33 (1996) 87–104 Dang, M., MILS-2 (2007) 473–500 D’Arcy, P. F., MIBS-14 (1982) 1–35 Darnall, D. W., MIBS-6 (1976) 251–290 Daron, H. H., MIBS-11 (1980) 337–376 Das, R., MIBS-27 (1991) 323–359 Dash, K. C., MIBS-14 (1982) 179–205 Datwyler, S. A., MIBS-38 (2001) 213–254 Daub, E., MILS-2 (2007) 445–471 Daune, M., MIBS-3 (1974) 1–43 Davidge, J., MIBS-41 (2004) 1–39 Davis, A. K., MILS-4 (2008) 255–294 Davis, J. M., MIBS-6 (1976) 291–321 Davydova, S. L., MIBS-8 (1979) 183–206 Dawson, A. A., MIBS-10 (1980) 129–166 Dawson, J. H., MILS-3 (2007) 319–359 Devez, A., MILS-5 (2009) 441–481 De Voss, J. J., MILS-3 (2007) 397–435 Dean, D. R., MIBS-39 (2002) 163–186 Debrunner, P. G., MIBS-7 (1978) 241–275 Debus, R. J., MIBS-37 (2000) 657–711 DeRose, E., MIBS-33 (1996) 619–648 Dikanov, S. A., MIBS-22 (1987) 207–263 Dillon, C. T., MIBS-41 (2004) 253–277 Djordjevic, C., MIBS-31 (1995) 595–616 Dobbek, H., MIBS-39 (2002) 227–263 Dodd, N. J. F., MIBS-10 (1980) 95–128 Dolderer, B., MILS-5 (2009) 83–104 Dolman, R. C., MIBS-42 (2004) 297–322 Dooley, D. M., MIBS-30 (1994) 361–403 Double, K. L., MILS-1 (2006) 125–149 Downey, K. M., MIBS-25 (1989) 1–30 Draganescu, A., MIBS-33 (1996) 453–484 Drakenberg, T., MIBS-25 (1989) 255–307 Driscoll, C. T., MIBS-24 (1988) 59–122 Dubler, E., MIBS-32 (1996) 301–338 Dukes, G. R., MIBS-1 (1974) 157–212 Durham, B., MIBS-27 (1991) 223–264

Met. Ions Life Sci. 2009, 6, 497–510

Durlach, J., MIBS-26 (1990) 549–578 Dwek, R. A., MIBS-4 (1974) 61–210

E Eady, R. R., MIBS-31 (1995) 363–405 Eason, P. D., MIBS-33 (1996) 427–452 Easwaran, K. R. K., MIBS-19 (1985) 109–137 Ebel, H., MIBS-26 (1990) 215–225; MIBS-26 (1990) 227–248 Ecker, D. J., MIBS-25 (1989) 147–169 Edwards, S. L., MIBS-25 (1989) 477–503 Egli, T. W., MIBS-28 (1992) 1–39 Eichenberger, E., MIBS-20 (1986) 67–100 Eichhorn, G. L., MIBS-10 (1980) 1–21 Eidsness, M. K., MIBS-27 (1991) 199–222 Einspahr, H., MIBS-17 (1984) 51–97 Elin, R. J., MIBS-26 (1990) 579–596 Emery, T., MIBS-7 (1978) 77–126 Enemark, J. H., MIBS-39 (2002) 621–654 English, A. M., MIBS-38 (2001) 313–350 Ensunsa, J. L., MIBS-37 (2000) 89–121 Epple, M., MILS-4 (2008) 207–217 Erdmann, F., MILS-2 (2007) 501–518 Ermler, U., MILS-6 (2009) 219–240 Ernst, F. D., MILS-2 (2007) 545–579 Evans, C. A., MIBS-9 (1979) 41–75; MIBS-9 (1979) 103–141 Eveleigh, D. E., MIBS-28 (1992) 399–414

F Farago, M. E., MIBS-23 (1988) 47–90 Faraone-Mennella, J., MILS-1 (2006) 9–60 Farkas, E., MILS-2 (2007) 63–107 Farrell, N., MIBS-32 (1996) 603–639; MIBS42 (2004) 251–296 Fauque, G. D., MIBS-23 (1988) 285–314 Fee, J. A., MIBS-13 (1981) 259–298 Feig, A. L., MIBS-37 (2000) 157–182 Feiters, M. C., MIBS-38 (2001) 461–655 Fenna, R. E., MIBS-30 (1994) 25–75 Fett, J. P., MIBS-35 (1998) 187–214 Fetzner, S., MIBS-39 (2002) 405–430; MIBS39 (2002) 481–537 Field, J. A., MIBS-28 (1992) 61–97 Fischer, B., MIBS-39 (2002) 265–316 Fischer, G., MILS-2 (2007) 501–518

AUTHOR INDEX

501

Fishel, L. A., MIBS-25 (1989) 477–503 Fiskum, G., MIBS-17 (1984) 187–214 Fitzgerald, W. F., MIBS-34 (1997) 53–111 Fleming, B. D., MILS-3 (2007) 127–155 Flessel, C. P., MIBS-10 (1980) 23–54 Follmann, H., MIBS-30 (1994) 131–161 Fontecilla-Camps, J. C., MILS-6 (2009) 151–178 Forseń, S., MIBS-25 (1989) 255–307 Forssell-Aronsson, E., MIBS-42 (2004) 77–108 Fortin, D., MILS-4 (2008) 377–411 Foster, M. A., MIBS-10 (1980) 129–166 Foulkes, E. C., MIBS-20 (1986) 157–200 Fox, M. R. S., MIBS-20 (1986) 201–228 Frank, P., MIBS-31 (1995) 423–490 Frankel, R. B., MILS-4 (2008) 343–376 Frankenberger Jr., W. T., MIBS-29 (1993) 185–227 Fratzl, P., MILS-4 (2008) 547–575 Frazzon, J., MIBS-39 (2002) 163–186 Freisinger, E., MILS-5 (2009) 107–153 Friberg, L. T., MIBS-34 (1997) 371–403 Fricker, S. P., MIBS-36 (1999) 665–721; MIBS-41 (2004) 421–480 Frieden, E., MIBS-13 (1981) 1–14; MIBS-13 (1981) 117–142 Friedman, M. E., MIBS-11 (1980) 337–376 Frøystein, N. A˚., MIBS-32 (1996) 397–418 Fry, C. H., MIBS-26 (1990) 463–488 Fu, P. K.-L., MIBS-40 (2003) 323–353 Fudge, R. J. P., MIBS-34 (1997) 259–287 Fukuto, J. M., MIBS-36 (1999) 547–595 Fung, H.-L., MIBS-36 (1999) 723–749 Furst, A., MIBS-10 (1980) 23–54

G Gajda-Schrantz, K., MILS-1 (2006) 371–393 Galanski, M., MIBS-42 (2004) 179–208 Galdes, A., MIBS-15 (1983) 1–54 Galka, M.; MIBS-38 (2001) 385–409 Galli, G., MIBS-28 (1992) 415–453 Galliot, M., MIBS-16 (1983) 245–260 Gampp, H., MIBS-12 (1981) 133–189; MIBS-22 (1987) 105–127 Garner, C. D., MIBS-31 (1995) 617–670; MIBS-39 (2002) 699–726 Ga¨rtner, W., MILS-2 (2007) 279–322 Gellert, R. W., MIBS-8 (1979) 1–55

Gelpke, M. D. S., MIBS-37 (2000) 559–586 Geraldes, C. F. G. C., MIBS-40 (2003) 513–588 Gergely, A., MIBS-9 (1979) 77–102; MIBS-9 (1979) 143–172 Gerlach, M., MILS-1 (2006) 125–149 Gibbs, E. J., MIBS-33 (1996) 367–397 Giedroc, D. P., MIBS-25 (1989) 171–234 Gillam, E. M. J., MILS-3 (2007) 477–560 Ginj, M., MIBS-42 (2004) 109–142 Girvan, H. M., MILS-3 (2007) 285–317 Gladyshev, V. N., MIBS-39 (2002) 655–672 Glasauer, S., MILS-4 (2008) 377–411 Gledhill, M., MILS-5 (2009) 441–481 Gmaj, P., MIBS-17 (1984) 99–127 Gold, M. H., MIBS-37 (2000) 559–586 Gormley, N. A., MIBS-37 (2000) 345–364 Gorren, A. C. F., MIBS-27 (1991) 199–222 Go¨tz, M. E., MILS-1 (2006) 125–149 Goyal, A., MIBS-28 (1992) 399–414 Graham, D. E., MILS-2 (2007) 357–415 Gralla, E. B., MIBS-36 (1999) 125–177 Grandi, G., MIBS-28 (1992) 415–453 Gra¨slund, A., MIBS-30 (1994) 109–129 Gravert, D. J., MIBS-33 (1996) 515–536 Gray, H. B, MIBS-27 (1991) 97–127; MILS1 (2006) 9–60; MILS-3 (2007) 157–185 Greger, J. L., MIBS-24 (1988) 199–215 Gresh, N., MIBS-19 (1985) 335–386 Griesser, R., MIBS-1 (1974) 213–247 Griffin, J. H., MIBS-33 (1996) 515–536 Grisham, C. M., MIBS-32 (1996) 1–26 Grubbs, R. D., MIBS-26 (1990) 177–192 Grundstro¨m, T., MIBS-25 (1989) 255–307 Grunes, D. L., MIBS-26 (1990) 33–56 Guengerich, F. P., MILS-3 (2007) 561–589 Guerinot, M. L., MIBS-35 (1998) 187–214 Guevremont, R., MIBS-9 (1979) 41–75; MIBS-9 (1979) 103–141 Guiet-Bara, A., MIBS-26 (1990) 549–578 Gunsalus, I. C., MIBS-7 (1978) 241–275 Gu¨nther, T., MIBS-26 (1990) 193–213; MIBS-26 (1990) 215–225

H Habash, J., MIBS-37 (2000) 279–304 Hajdu, J., MIBS-19 (1985) 53–80 Haley, B. E., MIBS-34 (1997) 461–478 Halford, S. E., MIBS-37 (2000) 345–364

Met. Ions Life Sci. 2009, 6, 497–510

502 Hall, B. D., MIBS-34 (1997) 259–287 Hall, M. D., MIBS-42 (2004) 297–322 Hall, S. K., MIBS-26 (1990) 463–488 Hamada, R., MIBS-34 (1997) 405–420 Hambley, T. W., MIBS-41 (2004) 253–277; MIBS-42 (2004) 297–322 Hammel, K. E., MIBS-28 (1992) 41–60 Hammond, P. B., MIBS-20 (1986) 157–200 Hancock Jr., A. B., MIBS-30 (1994) 163–199 Ha¨nsel, W., MIBS-14 (1982) 243–255 Hanson, P. J., MIBS-34 (1997) 185–212 Hantke, K., MIBS-35 (1998) 67–145 Harayama, S., MIBS-28 (1992) 99–156 Hare, L., MILS-5 (2009) 239–277 Harper, D. B., MIBS-29 (1993) 345–388 Harrison, P. M., MIBS-35 (1998) 435–477 Harrowfield, J. M., MIBS-40 (2003) 105–159 Hart, P. J., MILS-1 (2006) 179–205 Hartmanis, M. G. N., MIBS-30 (1994) 201–215 Hartmann, H.-J., MIBS-36 (1999) 389–413; MILS-5 (2009) 83–104 Harvey, C. F., MIBS-44 (2005) 145–169 Hatano, M., MIBS-5 (1976) 245–277 Hauenstein Jr., B. L., MIBS-13 (1981) 319–347 Hausinger, R. P., MILS-2 (2007) 519–544 Hay, R. W., MIBS-5 (1976) 127–172; MIBS5 (1976) 173–243 Hayes, R. L., MIBS-16 (1983) 279–315 Heaton, F. W., MIBS-26 (1990) 119–133 Heck, J. D., MIBS-20 (1986) 259–278; MIBS-20 (1986) 279–303 Hefford, R. J. W., MIBS-9 (1979) 173–212 Heller, M. J., MIBS-1 (1974) 1–49 Helliwell, J. R., MIBS-37 (2000) 279–304 Helm, L., MIBS-40 (2003) 589–641 Hempstead, P. D., MIBS-35 (1998) 435–477 Herzfeld, J., MIBS-7 (1978) 311–349 Heydorn, K., MIBS-16 (1983) 123–138; MIBS-16 (1983) 167–183; MIBS-16 (1983) 225–234 Heyduk, E., MIBS-38 (2001) 255–287 Heyduk, T., MIBS-38 (2001) 255–287 Hidai, M., MIBS-39 (2002) 121–161 Hidalgo, J., MILS-5 (2009) 279–317 Hider, R. C., MIBS-35 (1998) 691–730 Hildebrand, M., MILS-4 (2008) 255–294 Hill, H. A. O., MIBS-27 (1991) 431–494 Hille, R., MIBS-39 (2002) 187–226; MILS-6 (2009) 395–416

Met. Ions Life Sci. 2009, 6, 497–510

AUTHOR INDEX Hinton, J. F., MIBS-19 (1985) 173–206 Hobman, J. L., MIBS-34 (1997) 527–568 Hodgson, K. O., MIBS-31 (1995) 423–490 Hoehler, T. M., MIBS-43 (2005) 9–48 Hoffelt, J., MIBS-16 (1983) 245–260 Hoganson, C. W., MIBS-30 (1994) 77–107; MIBS-37 (2000) 613–656 Hom, P., MIBS-26 (1990) 285–320 Honek, J. F., MILS-2 (2007) 445–471 Ho¨rmann, H., MIBS-3 (1974) 89–132 Horrocks Jr., W. DeW., MIBS-40 (2003) 281–322 Hoskins, N., MILS-3 (2007) 437–476 Howe-Grant, M. E., MIBS-11 (1980) 63–125 Hsieh, W.-Y., MIBS-37 (2000) 429–504 Huang, X., MIBS-36 (1999) 309–364 Huber, R., MIBS-39 (2002) 227–263 Hu¨bner, K. F., MIBS-16 (1983) 279–315 Huffman, D. L., MIBS-34 (1997) 503–526 Hughes, E. R., MIBS-7 (1978) 351–376 Huguet, J., MIBS-15 (1983) 55–99 Hultman, P., MIBS-34 (1997) 421–440 Hunter, D. J. B., MILS-3 (2007) 477–560 Hunter, N. S., MIBS-37 (2000) 279–304 Hunter, R. C., MILS-4 (2008) 127–165 Hunziker, N., MIBS-26 (1990) 531–547 Hu¨ttermann, J., MIBS-22 (1987) 1–80; MIBS-37 (2000) 365–405; MIBS-39 (2002) 481–537 Hwang, J., MIBS-37 (2000) 527–557 Hydes, P. C., MIBS-11 (1980) 1–62

I Iitaka, Y., MIBS-19 (1985) 207–227 Inesi, G., MIBS-17 (1984) 129–186 Ishii, T., MIBS-31 (1995) 491–509 Isied, S. S., MIBS-27 (1991) 1–56 Issaq, H. J., MIBS-10 (1980) 55–93

J Jack, T. R., MIBS-8 (1979) 159–182 Jacobs, R. M., MIBS-20 (1986) 201–228 Jakupec, M. A., MIBS-42 (2004) 179–208; MIBS-42 (2004) 425–462 Jameson, G. N. L., MILS-1 (2006) 281–320 Jameson, R. F., MIBS-12 (1981) 1–30; MILS-1 (2006) 281–320

AUTHOR INDEX

503

Janssen, D. B., MIBS-28 (1992) 299–327 Jansson, B., MIBS-10 (1980) 281–311 Jarnum, S., MIBS-15 (1983) 415–438 Jarrell, W. M., MIBS-20 (1986) 119–156 Jaun, B., MIBS-29 (1993) 287–337; MILS-2 (2007) 323–356; MILS-6 (2009) 115–132 Jellinger, K. A., MILS-1 (2006) 281–320 Jensen, A., MIBS-16 (1983) 139–150; MIBS-16 (1983) 151–165; MIBS-16 (1983) 167–183; MIBS-16 (1983) 185–199; MIBS-16 (1983) 201–212; MIBS-16 (1983) 213–223; MIBS-16 (1983) 225–234; MIBS-16 (1983) 235–243; MIBS-18 (1984) 5–59; MIBS-18 (1984) 61–103 Jin, S., MILS-3 (2007) 319–359 Joesten, M. D., MIBS-11 (1980) 285–304 Johansson, C., MIBS-25 (1989) 255–307 Johnston, D. H., MIBS-33 (1996) 297–324 Jokinen, R., MIBS-26 (1990) 15–32 Jones, M. M., MIBS-16 (1983) 47–83 Jones, P. W., MIBS-41 (2004) 139–183 Jones, P., MIBS-7 (1978) 185–240 Jøns, O., MIBS-16 (1983) 201–212 Jørgensen, S. E., MIBS-18 (1984) 5–59; MIBS-18 (1984) 61–103 Ju, T., MILS-2 (2007) 473–500 Jung, C., MILS-3 (2007) 187–234

K Kaden, T. A., MIBS-4 (1974) 1–27 Ka¨gi, J. H. R., MIBS-15 (1983) 213–273 Kaiser, E. T., MIBS-25 (1989) 395–415 Kalb (Gilboa), R. J., MIBS-37 (2000) 279–304 Kalbitzer, H. R., MIBS-22 (1987) 81–103 Kanamori, K., MIBS-31 (1995) 45–88 Kanopkaite˙ , S., MIBS-10 (1980) 253–279 Kappl, R., MIBS-22 (1987) 1–80; MIBS-37 (2000) 365–405; MIBS-39 (2002) 481–537 Karlin, K. D., MILS-6 (2009) 295–361 Karlson, U., MIBS-29 (1993) 185–227 Kartha, S., MIBS-27 (1991) 323–359 Kasprzak, K. S., MILS-2 (2007) 581–618; MILS-2 (2007) 619–660 Keen, C. L., MIBS-37 (2000) 89–121 Kelley, S. O., MIBS-36 (1999) 211–249

Kendall, D. A., MIBS-25 (1989) 395–415 Kennedy, B. J., MIBS-41 (2004) 253–277 Kennedy, M. C., MIBS-32 (1996) 579–602 Keppler, B. K., MIBS-42 (2004) 179–208; MIBS-42 (2004) 425–462 Kerr, D. N. S., MIBS-24 (1988) 217–258 Khutsishvili, I. G., MIBS-33 (1996) 253–267 Killian, C. E., MILS-4 (2008) 37–69 Kim, J. E., MILS-1 (2006) 9–60 Kim, J. K., MILS-2 (2007) 519–544 Kim, K.-H., MIBS-34 (1997) 185–212 Kimura, E., MIBS-33 (1996) 29–52 Kirchgessner, M., MIBS-15 (1983) 319–361; MIBS-15 (1983) 363–414 Kiss, T., MIBS-9 (1979) 143–172; MILS-1 (2006) 371–393 Kist, M., MILS-2 (2007) 545–579 Kito, M., MIBS-38 (2001) 187–196 Klevickis, C., MIBS-32 (1996) 1–26 Klimmek, O., MIBS-43 (2005) 105–130 Knowles, P. F., MIBS-30 (1994) 361–403 Kodadek, T., MIBS-38 (2001) 351–384 Koenig, S. H., MIBS-21 (1987) 229–270 Koeppe II, R. E., MIBS-19 (1985) 173–206 Komiyama, M., MIBS-38 (2001) 25–41; MIBS-40 (2003) 355–368; MIBS-40 (2003) 463–475 Koppenol, W. H., MIBS-36 (1999) 597–619 Ko¨ster, W., MIBS-35 (1998) 67–145 Kostic´, N. M., MIBS-27 (1991) 129–182; MIBS-38 (2001) 145–186 Kowalik-Jankowska, T., MILS-2 (2007) 63–107 Kozarich, J. W., MIBS-30 (1994) 279–313 Kozelka, J., MIBS-33 (1996) 1–28 Kozlowski, H., MILS-1 (2006) 61–87; MILS-2 (2007) 63–107 Kraatz, H.-B., MIBS-38 (2001) 385–409 Krachler, M., MIBS-44 (2005) 171–203 Kraut, J., MIBS-25 (1989) 477–503 Kraütler, B., MILS-6 (2009) 1–51 Krezoski, S., MILS-5 (2009) 353–397 Kroneck, P. M. H., MIBS-28 (1992) 455–505; MIBS-39 (2002) 369–403; MIBS-43 (2005) 1–7; MIBS-43 (2005) 75–103; MIBS-44 (2005) 97–144; MILS-2 (2007) 31–62; MILS-6 (2009) 363–393 Kruck, T. P. A., MIBS-24 (1988) 285–314 Kuo, L. Y., MIBS-33 (1996) 53–85

Met. Ions Life Sci. 2009, 6, 497–510

504

AUTHOR INDEX

Ku¨pper, H., MIBS-44 (2005) 97–144; MILS2 (2007) 31–62 Kusters, J. G., MILS-2 (2007) 545–579 Kustin, K., MIBS-31 (1995) 511–542 Kuusela, S., MIBS-32 (1996) 271–300

L Ladefoged, K., MIBS-15 (1983) 415–438 Lagarde, F., MIBS-39 (2002) 741–759 Lai, J. C. K., MIBS-37 (2000) 123–156 Landgraf, R., MIBS-33 (1996) 485–513 Landolph, J. R., MIBS-36 (1999) 445–483 Langley, S., MILS-4 (2008) 377–411 Lappin, G., MIBS-13 (1981) 15–71 Larsson, L., MIBS-41 (2004) 71–102 Lawson, D. M., MIBS-39 (2002) 31–74; MIBS-39 (2002) 75–119 Lay, P. A., MIBS-41 (2004) 253–277 Le Roux, G., MIBS-43 (2005) 239–275 Lean, D. R. S., MIBS-43 (2005) 221–238 Lee, B. T. O., MIBS-30 (1994) 405–434 Lee, D. B. N., MIBS-26 (1990) 285–320 Lee, J. C., MILS-1 (2006) 9–60 Lee, Y.-H., MIBS-34 (1997) 113–130 LeGall, J., MIBS-23 (1988) 285–314 Leng, M., MIBS-33 (1996) 87–104 Leónard, A., MIBS-20 (1986) 229–258 Leong, S. A., MIBS-35 (1998) 147–186 Lerch, K., MIBS-13 (1981) 143–186; MIBS13 (1981) 299–318 Leroy, M., MIBS-39 (2002) 741–759 Lettinga, G., MIBS-28 (1992) 61–97 Leussing, D. L., MIBS-2 (1973) 127–166; MIBS-5 (1976) 1–77 LeVier, K., MIBS-35 (1998) 187–214 Levitin, I. Y., MIBS-36 (1999) 485–519 Lewis, B. L., MIBS-29 (1993) 79–99 Li, R., MIBS-40 (2003) 707–751 Li, W., MIBS-33 (1996) 619–648 Liang, Q., MIBS-33 (1996) 427–452 Libman, J., MIBS-35 (1998) 329–354 Lichtenegger, H. C., MILS-4 (2008) 295–325 Liebert, C. A., MIBS-34 (1997) 441–460 Lillig, C. H., MILS-5 (2009) 413–439 Lim, L., MIBS-37 (2000) 123–156 Lincoln, P., MIBS-33 (1996) 177–252 Lindahl, P. A., MILS-2 (2007) 357–415; MILS-6 (2009) 133–150 Lindberg, S. E., MIBS-34 (1997) 185–212

Met. Ions Life Sci. 2009, 6, 497–510

Linder, P. W., MIBS-7 (1978) 29–76 Linert, W., MILS-1 (2006) 281–320 Linse, S., MIBS-25 (1989) 255–307 Liochev, S. I., MIBS-36 (1999) 1–39 Lippard, S. J., MIBS-11 (1980) 63–125; MIBS-32 (1996) 687–726; MIBS-42 (2004) 143–177; MILS-1 (2006) 321–370 Lippert, B., MIBS-33 (1996) 105–141; MIBS33 (1996) 143–176 Lipscomb, J. D., MIBS-28 (1992) 243–298 Liptak, M. D., MILS-6 (2009) 417–460 Liu, A. H., MIBS-33 (1996) 53–85 Liu, J., MILS-5 (2009) 399–412 Liu, X. S., MILS-4 (2008) 327–341 Lloyd, J. R, MIBS-44 (2005) 205–240 Lobreáux, S., MIBS-35 (1998) 563–584 Long, E. C., MIBS-33 (1996) 427–452 Lo¨nnberg, H., MIBS-32 (1996) 271–300 Lontie, R., MIBS-3 (1974) 183–200; MIBS13 (1981) 229–258 Lowe, D. J., MIBS-39 (2002) 455–479 Lu, P. H., MILS-1 (2006) 151–177 Lu, Y., MILS-3 (2007) 267–284 Lubitz, W., MILS-2 (2007) 279–322 Lucas, H. R., MILS-6 (2009) 295–361 Luchinat, C., MIBS-15 (1983) 101–156; MIBS-21 (1987) 47–86; MIBS-40 (2003) 513–588 Luczkowski, M., MILS-1 (2006) 61–87 Ludwig, B., MIBS-44 (2005) 75–96 Lyons, T. J., MIBS-36 (1999) 125–177

M Mabbs, F. E., MIBS-31 (1995) 617–670 Maecke, H. R., MIBS-42 (2004) 109–142 Magnusson, P., MIBS-41 (2004) 71–102 Magos, L., MIBS-34 (1997) 321–370 Maguire, M. E., MIBS-26 (1990) 135–153 Makinen, M. W., MIBS-22 (1987) 129–206; MIBS-31 (1995) 89–127 Makrigiorgos, G. M., MIBS-36 (1999) 521–545 Malcolm, E. G., MIBS-43 (2005) 195–219 Malnoe¨, A., MIBS-17 (1984) 215–273 Mao, Q., MIBS-33 (1996) 619–648 Marafante, E., MIBS-29 (1993) 161–184 Marcon, G., MIBS-41 (2004) 279–304; MIBS-42 (2004) 385–424

AUTHOR INDEX Margerum, D. W., MIBS-1 (1974) 157–212; MIBS-12 (1981) 75–132 Mariam, Y. H., MIBS-8 (1979) 57–124 Marier, J. R., MIBS-26 (1990) 85–104 Marin, F., MILS-4 (2008) 71–126 Marks, T. J., MIBS-33 (1996) 53–85 Marnett, L. J., MIBS-30 (1994) 163–199 Maroney, M. J., MIBS-21 (1987) 87–120; MILS-2 (2007) 417–443 Martell, A. E., MIBS-2 (1973) 207–268; MIBS-16 (1983) 85–102 Martin, J. D., MIBS-44 (2005) 21–46 Martin, L. L., MILS-3 (2007) 127–155 Martin, R. B., MIBS-1 (1974) 129–156; MIBS-8 (1979) 57–124; MIBS-9 (1979) 1–39; MIBS-17 (1984) 1–49; MIBS-19 (1985) 19–52; MIBS-20 (1986) 21–65; MIBS-23 (1988) 123–164; MIBS-23 (1988) 315–330; MIBS-24 (1988) 1–57; MIBS-26 (1990) 1–13; MIBS-32 (1996) 61–89; MIBS-38 (2001) 1–23 Martin, R. P., MIBS-2 (1973) 1–61 Mason, R. P., MIBS-34 (1997) 53–111 Masters, C. L., MIBS-36 (1999) 365–387 Masuda, H., MIBS-32 (1996) 207–270 Matsumoto, K., MIBS-40 (2003) 191–232 Matthews, R. G., MILS-6 (2009) 53–114 Matzapetakis, M., MILS-4 (2008) 327–341 Mauro, J. M., MIBS-25 (1989) 477–503 May, P. M., MIBS-7 (1978) 29–76; MIBS-12 (1981) 283–317 Mayer, P. H., MIBS-29 (1993) 79–99 Mayland, H. F., MIBS-26 (1990) 33–56 McCormick, D. B., MIBS-1 (1974) 213–247 McElnay, J. C., MIBS-14 (1982) 1–35 McLachlan, D. R., MIBS-24 (1988) 285–314 McLean, K. J., MILS-3 (2007) 285–317 McLendon, G., MIBS-27 (1991) 183–198 McLennan, S. M., MIBS-40 (2003) 1–38 McMillin, D. R., MIBS-13 (1981) 319–347 McNeill, J. H., MIBS-31 (1995) 575–594 Meade, T. J., MIBS-32 (1996) 453–478; MIBS-42 (2004) 1–38 Meares, C. F., MIBS-38 (2001) 213–254 Meharenna, Y. T., MILS-3 (2007) 57–96 Meier, R., MIBS-31 (1995) 45–88 Meili, M., MIBS-34 (1997) 21–51 Melius, P., MIBS-11 (1980) 337–376 Meloni, G., MILS-5 (2009) 319–351 Mendel, R. R., MIBS-39 (2002) 317–368 Mennie, D., MIBS-29 (1993) 37–77

505 Menon, C. R., MIBS-23 (1988) 359–402 Merbach, A. E., MIBS-40 (2003) 589–641 Messori, L., MIBS-21 (1987) 47–86; MIBS-41 (2004) 279–304; MIBS-42 (2004) 385–424 Mestroni, G., MIBS-42 (2004) 323–351 Meunier, B., MIBS-33 (1996) 399–426 Meyer, F., MILS-2 (2007) 181–239 Meyer, H., MIBS-26 (1990) 57–83 Meyerstein, D., MIBS-36 (1999) 41–77 Michel, K., MIBS-44 (2005) 75–96 Mikkelsen, K. V., MIBS-27 (1991) 57–96 Mildvan, A. S., MIBS-25 (1989) 309–334 Miller Frey, C., MIBS-1 (1974) 51–116 Miller, M. A., MIBS-25 (1989) 477–503 Millett, F., MIBS-27 (1991) 223–264 Milovic´, N. M., MIBS-38 (2001) 145–186 Minski, M. J., MIBS-37 (2000) 123–156 Mitzenheim, S., MIBS-31 (1995) 45–88 Mizobe, Y., MIBS-39 (2002) 121–161 Moir, R. D., MIBS-36 (1999) 309–364 Monaselidze, J. R., MIBS-23 (1988) 331–357 Monje, P. V., MILS-4 (2008) 219–254 Montfort, W. R., MIBS-36 (1999) 621–663 Moore, G. R., MIBS-21 (1987) 187–227 Moradian-Oldak, J., MILS-4 (2008) 507–546 Morel, F. M. M., MIBS-35 (1998) 1–36; MIBS-43 (2005) 195–219 Morel, I., MIBS-36 (1999) 251–287 Morgan, J. J., MIBS-37 (2000) 1–34 Mori, S., MIBS-35 (1998) 215–238 Morris, P. J., MIBS-5 (1976) 173–243 Morrison, H., MIBS-33 (1996) 269–296 Morrow, J. R., MIBS-33 (1996) 561–592 Mosulishvili, L. M., MIBS-10 (1980) 167–206 Mottet, N. K., MIBS-34 (1997) 371–403 Moura, I., MIBS-23 (1988) 285–314 Moura, J. J. G., MIBS-23 (1988) 285–314; MIBS-39 (2002) 539–570 Muhlrad, A., MIBS-31 (1995) 211–230 Mulazzani, Q. G., MIBS-36 (1999) 103–123 Mulrooney, S. B., MILS-2 (2007) 519–544 Multhaup, G., MIBS-36 (1999) 365–387; MILS-1 (2006) 115–123 Munro, A. W., MILS-3 (2007) 285–317 Murer, H., MIBS-17 (1984) 99–127 Murphy, W. L., MILS-4 (2008) 577–606 Mustafi, D., MIBS-31 (1995) 89–127

Met. Ions Life Sci. 2009, 6, 497–510

506

AUTHOR INDEX

N Nakai, I., MIBS-31 (1995) 491–509 Nakanishi, K., MIBS-31 (1995) 423–490 Nancollas, G. H., MILS-4 (2008) 413–456 Natile, G., MIBS-42 (2004) 209–250 Navon, G., MIBS-21 (1987) 1–45 Nawata, Y., MIBS-19 (1985) 207–227 Neilands, J. B., MIBS-19 (1985) 313–333 Nelsestuen, G. L., MIBS-17 (1984) 353–380 Nerdal, W., MIBS-34 (1997) 479–501 Nieboer, E., MIBS-23 (1988) 91–121; MIBS-23 (1988) 359–402 Nielsen, F. H., MIBS-31 (1995) 543–573 Nieminen, T. M., MILS-2 (2007) 1–29 Nishino, T., MIBS-39 (2002) 431–454 Nohl, H., MIBS-36 (1999) 289–307 Nordberg, G. F., MILS-5 (2009) 1–29 Nordberg, M., MILS-5 (2009) 1–29 Nordeń, B., MIBS-33 (1996) 177–252 Norris, J. R., MIBS-27 (1991) 323–359 Nozawa, T., MIBS-5 (1976) 245–277 Nriagu, J. O., MIBS-34 (1997) 131–160

O O’Brien, P., MIBS-19 (1985) 295–312 Ochiai, E.-I., MIBS-30 (1994) 1–24; MIBS-30 (1994) 255–278 Odani, A., MIBS-32 (1996) 207–270 O’Driscoll, N. J., MIBS-43 (2005) 221–238 O’Halloran, K., MIBS-35 (1998) 667–690 O’Halloran, T. V., MIBS-25 (1989) 105–146; MIBS-32 (1996) 557–578; MIBS-34 (1997) 503–526 Okoshi, K., MIBS-31 (1995) 491–509 Onuchic, J. N., MIBS-27 (1991) 97–127 Orme-Johnson, W. H., MIBS-7 (1978) 127–183 Orvig, C., MIBS-31 (1995) 575–594; MIBS-41 (2004) 221–252 Orville, A. M., MIBS-28 (1992) 243–298 Osame, M., MIBS-34 (1997) 405–420 Osinsky, S. P., MIBS-36 (1999) 485–519 O¨sterberg, R., MIBS-3 (1974) 45–88 Otsuka, M., MIBS-32 (1996) 27–60 OuYang, B., MILS-2 (2007) 473–500 Owens, G. D., MIBS-12 (1981) 75–132

Met. Ions Life Sci. 2009, 6, 497–510

P Pagani, G. M., MILS-2 (2007) 473–500 Page, A. L., MIBS-18 (1984) 287–332 Pagenkopf, G. K., MIBS-20 (1986) 101–118 Pai, E. F., MIBS-39 (2002) 431–454 Paine, M. L., MILS-4 (2008) 507–546 Painter, G. R., MIBS-19 (1985) 229–294 Paquette, S. J., MIBS-40 (2003) 69–104 Parker, D., MIBS-40 (2003) 233–280 Parkin, G., MIBS-38 (2001) 411–460 Pasternack, R. F., MIBS-33 (1996) 367–397 Paterson, M. J., MIBS-34 (1997) 259–287 Pau, R. N., MIBS-39 (2002) 31–74 Paytan, A., MIBS-43 (2005) 153–193 Pearson, L., MIBS-33 (1996) 485–513 Pecoraro, V. L., MIBS-37 (2000) 429–504 Pendergrass, J. C., MIBS-34 (1997) 461–478 Penkowa, M., MILS-5 (2009) 279–317 Penner-Hahn, J. E., MIBS-37 (2000) 527–557 Perera, R., MILS-3 (2007) 319–359 Perl, D. P., MIBS-24 (1988) 259–283 Perlman, S., MILS-1 (2006) 151–177 Perrin, D. D., MIBS-2 (1973) 167–206; MIBS-14 (1982) 207–241 Perrin, D. M., MIBS-33 (1996) 485–513 Persson, P., MIBS-16 (1983) 167–183; MIBS-16 (1983) 185–199; MIBS-16 (1983) 201–212; MIBS-16 (1983) 213–223; MIBS-16 (1983) 225–234 Peryea, F. J., MIBS-20 (1986) 119–156 Petering, D. H., MIBS-11 (1980) 197–229; MIBS-33 (1996) 619–648; MIBS-42 (2004) 463–497; MILS-5 (2009) 353–397 Peters, J. W., MILS-6 (2009) 179–218 Petit-Ramel, M. M., MIBS-2 (1973) 1–61 Pettit, L. D., MIBS-9 (1979) 173–212 Pfister, T. D., MILS-3 (2007) 267–284 Pletneva, E. V., MILS-1 (2006) 9–60 Pochapsky, T. C., MILS-2 (2007) 473–500 Pocker, Y., MIBS-25 (1989) 335–358 Pocklington, T., MIBS-10 (1980) 129–166 Pohl, H. R., MILS-1 (2006) 395–425 Pollard, K. M., MIBS-34 (1997) 421–440 Polzin, G. M., MIBS-38 (2001) 103–143 Pories, W. J., MIBS-10 (1980) 207–251 Potter, J. D., MIBS-17 (1984) 381–410 Poulos, T. L., MIBS-30 (1994) 25–75; MILS-3 (2007) 57–96 Powell, A. K., MIBS-35 (1998) 515–561

AUTHOR INDEX

507

Poyner, R. R., MIBS-37 (2000) 183–207 Prasad, A. S., MIBS-14 (1982) 37–55; MIBS-41 (2004) 103–137 Pratt, J. M., MIBS-29 (1993) 229–286 Pratviel, G., MIBS-33 (1996) 399–426 Pressman, B. C., MIBS-19 (1985) 1–18; MIBS-19 (1985) 229–294 Price, H. J., MIBS-37 (2000) 279–304 Price, N. M., MIBS-35 (1998) 1–36 Pullman, A., MIBS-19 (1985) 335–386 Pyle, A. M., MIBS-32 (1996) 479–520; MIBS-40 (2003) 477–512

Q Que Jr., L., MIBS-21 (1987) 87–120; MIBS-37 (2000) 505–525 Quiroz, S., MILS-2 (2007) 519–544

R Rabenstein, D. L., MIBS-9 (1979) 41–75; MIBS-9 (1979) 103–141 Radding, S. B., MIBS-10 (1980) 23–54 Raftery, J., MIBS-37 (2000) 279–304 Rajan, K. S., MIBS-6 (1976) 291–321 Rau, T., MIBS-32 (1996) 339–378 Rausch, N., MILS-2 (2007) 1–29 Rayssiguier, Y., MIBS-26 (1990) 341–358 Reed, C. A., MIBS-7 (1978) 277–310 Reed, G. H., MIBS-37 (2000) 183–207 Reedijk, J., MIBS-32 (1996) 641–685 Rehder, D., MIBS-31 (1995) 1–43 Rencz, A., MIBS-43 (2005) 221–238 Rengel, Z., MIBS-37 (2000) 57–87 Renshaw, J. C., MIBS-44 (2005) 205–240 Reynolds, M. F., MIBS-37 (2000) 505–525 Ribeiro, J. M. C., MIBS-36 (1999) 621–663 Riber, E., MIBS-16 (1983) 151–165; MIBS-16 (1983) 167–183; MIBS-16 (1983) 213–223; MIBS-16 (1983) 225–234 Ribeyre, F., MIBS-34 (1997) 289–319 Rickaby, R. E. M., MIBS-44 (2005) 241–268 Riederer, P., MILS-1 (2006) 125–149 Rifkind, J. M., MIBS-12 (1981) 191–232; MIBS-15 (1983) 275–317 Ringel, I., MIBS-31 (1995) 211–230 Riordan, J. F., MIBS-25 (1989) 359–394

Risher, J. F., MILS-1 (2006) 395–425 Rivera, M., MILS-6 (2009) 241–293 Robinson, C., MIBS-26 (1990) 489–504 Robinson, H. A., MIBS-23 (1988) 1–29 Robinson, M. F., MIBS-16 (1983) 1–26 Robinson, W. E., MIBS-31 (1995) 511–542 Rodrı´ guez, J. C., MILS-6 (2009) 241–293 Rokita, S. E., MIBS-33 (1996) 537–560; MIBS-38 (2001) 289–311 Romaõ, M. J., MIBS-39 (2002) 539–570 Ro¨sch, F., MIBS-42 (2004) 77–108 Rosen, B. P., MIBS-17 (1984) 129–186 Rosenberg, B., MIBS-11 (1980) 127–196 Rosenberg, D. M., MIBS-34 (1997) 259–287 Rossetto, F. E., MIBS-23 (1988) 359–402 Rossi, M., MIBS-42 (2004) 353–384 Roth, H.-P., MIBS-15 (1983) 363–414 Roy, A. T., MIBS-26 (1990) 285–320 Roy, R., MIBS-39 (2002) 673–697 Rubin, M., MIBS-16 (1983) 85–102 Rudd, J. W. M., MIBS-34 (1997) 259–287 Rudolf, M., MIBS-43 (2005) 75–103 Rusnak, F., MIBS-37 (2000) 305–343 Ryan, D. E., MIBS-31 (1995) 423–490 Ryan, M. P., MIBS-26 (1990) 249–269

S Sabat, M., MIBS-32 (1996) 521–555; MIBS-33 (1996) 143–176 Sagripanti, J.-L., MIBS-36 (1999) 179–209 Saji, H., MIBS-10 (1980) 313–340 Salnikow, K., MILS-2 (2007) 581–618; MILS-2 (2007) 619–660 Sanchez-Delgado, R. A., MIBS-41 (2004) 379–419 Sanford, W. E., MIBS-23 (1988) 91–121 Sarafian, T. A., MIBS-36 (1999) 415–444 Sarkar, B., MIBS-12 (1981) 233–281; MILS-1 (2006) 207–225 Satterlee, J. D., MIBS-21 (1987) 121–185 Sava, G., MIBS-42 (2004) 323–351 Savory, J., MIBS-24 (1988) 315–345; MIBS-24 (1988) 347–371 Schaffner, W., MILS-5 (2009) 31–49 Schanke, C. A., MIBS-28 (1992) 329–356 Scharff, J. P., MIBS-2 (1973) 1–61 Schecher, W. D., MIBS-24 (1988) 59–122 Schimatschek, H. F., MIBS-41 (2004) 41–69 Schindler, P. W., MIBS-18 (1984) 105–135

Met. Ions Life Sci. 2009, 6, 497–510

508 Schink, B., MIBS-43 (2005) 131–151 Schlichting, I., MILS-3 (2007) 235–265 Schmidbaur, H., MIBS-14 (1982) 179–205 Schneider, H.-J., MIBS-40 (2003) 369–462 Schneider, P. W., MIBS-6 (1976) 197–249 Schrag, D. P., MIBS-44 (2005) 241–268 Schu¨bbe, S., MILS-4 (2008) 343–376 Schuler, M. A., MILS-3 (2007) 1–26 Schumacher, W., MIBS-28 (1992) 455–505 Schwarz, G., MIBS-39 (2002) 317–368 Scott, R. A., MIBS-27 (1991) 199–222 Scozzafava, A., MIBS-12 (1981) 31–74 Seiler, H. G., MIBS-16 (1983) 317–353; MIBS-20 (1986) 305–336; MIBS-23 (1988) 403–428; MIBS-26 (1990) 611–624; MIBS-31 (1995) 671–688 Sellers, P., MIBS-25 (1989) 255–307 Sergent, O., MIBS-36 (1999) 251–287 Serpersu, E. H., MIBS-25 (1989) 309–334 Shanzer, A., MIBS-35 (1998) 329–354 Sharma, V. S., MIBS-2 (1973) 127–166 Shima, S., MILS-6 (2009) 219–240 Shionoya, M., MIBS-33 (1996) 29–52 Shotyk, W., MIBS-43 (2005) 239–275; MIBS-44 (2005) 171–203; MILS-2 (2007) 1–29 Sievers, C., MIBS-36 (1999) 389–413 Sigel, A., MIBS-29 (1993) 339–344 Sigel, H., MIBS-1 (1974) 213–247; MIBS-2 (1973) 63–125; MIBS-8 (1979) 125–158; MIBS-29 (1993) 339–344; MIBS-32 (1996) 135–205; MIBS-32 (1996) 207–270; MILS-2 (2007) 109–180 Sigel, R. K. O., MIBS-40 (2003) 477–512; MILS-2 (2007) 109–180 Sigg, L., MIBS-44 (2005) 47–73 Sigman, D. S., MIBS-33 (1996) 485–513 Silber, H. B., MIBS-40 (2003) 69–104 Silver, S., MIBS-30 (1994) 405–434 Sleep, N. H., MIBS-43 (2005) 49–73 Sletten, E., MIBS-32 (1996) 397–418; MIBS-34 (1997) 479–501 Sligar, S. G., MIBS-7 (1978) 241–275; MIBS25 (1989) 417–475; MILS-3 (2007) 1–26 Smith, B. E., MIBS-39 (2002) 75–119 Smith, M. J., MIBS-31 (1995) 423–490 Smith, T., MIBS-34 (1997) 441–460 Smith, W. L., MIBS-30 (1994) 163–199 Snavely, M. D., MIBS-26 (1990) 155–175 Snyder, M. J., MILS-3 (2007) 97–126 So, A. G., MIBS-25 (1989) 1–30

Met. Ions Life Sci. 2009, 6, 497–510

AUTHOR INDEX Solaro, R. J., MIBS-6 (1976) 323–398 Song, B., MIBS-32 (1996) 135–205 Sono, M., MILS-3 (2007) 319–359 Sorenson, J. R. J., MIBS-14 (1982) 77–124 Sosa-Torres, M. E., MILS-6 (2009) 363–393 So´va´go´, I., MIBS-9 (1979) 77–102; MILS-2 (2007) 63–107 Spa¨tling, L., MIBS-26 (1990) 513–529 Spence, J. T., MIBS-5 (1976) 279–324 Sposito, G., MIBS-18 (1984) 287–332; MIBS-20 (1986) 1–20 Springer, B. A., MIBS-25 (1989) 417–475 Stankiewicz, P. J., MIBS-31 (1995) 249–285; MIBS-31 (1995) 287–324 Stanley, H. E., MIBS-7 (1978) 311–349 Stayton, P. S., MIBS-25 (1989) 417–475 Stein, E. A., MIBS-17 (1984) 215–273 Steinnes, E., MIBS-44 (2005) 1–19 Steinrauf, L. K., MIBS-19 (1985) 139–171 Stemp, E. D. A., MIBS-33 (1996) 325–365 Stewart, L. J., MIBS-39 (2002) 699–726 Stiefel, E. I., MIBS-39 (2002) 1–29 St. Louis, V. L., MIBS-34 (1997) 259–287 Stokes, P., MIBS-23 (1988) 31–46 Stolze, K., MIBS-36 (1999) 289–307 Street, J. H., MIBS-43 (2005) 153–193 Strozyk, D., MILS-1 (2006) 1–7; MILS-1 (2006) 427–435 Stubbs, M., MIBS-32 (1996) 727–780 Stuehr, J., MIBS-1 (1974) 51–116 Stu¨nzi, H., MIBS-14 (1982) 207–241 Stu¨rzenbaum, S. R., MILS-5 (2009) 183–197 Sua´rez, L., MIBS-41 (2004) 379–419 Suarez-Gonzalez, D., MILS-4 (2008) 577–606 Suelter, C. H., MIBS-3 (1974) 201–251 Sugiura, Y., MIBS-19 (1985) 81–108 Suhy, D. A., MIBS-32 (1996) 557–578 Sukdeo, N., MILS-2 (2007) 445–471 Summers, A. O., MIBS-34 (1997) 441–460 Sun, H., MIBS-41 (2004) 333–378 Supkowski, R. M., MIBS-40 (2003) 281–322 Swinehart, J. H., MIBS-6 (1976) 141–196 Sykes, A. G., MIBS-27 (1991) 291–321 Szeto, K.-Y., MIBS-41 (2004) 333–378

T Tabatabai, N. M., MILS-5 (2009) 353–397 Tabrizi-Fard, M., MIBS-36 (1999) 723–749

AUTHOR INDEX

509

Takarada, T., MIBS-40 (2003) 355–368 Takita, T., MIBS-19 (1985) 81–108 Tanzi, R. E., MIBS-36 (1999) 309–364 Taylor, G. J., MIBS-24 (1988) 123–163; MIBS-24 (1988) 165–198 Taylor, S. R., MIBS-40 (2003) 1–38 Teixeira, M., MIBS-23 (1988) 285–314 Terreno, E., MIBS-40 (2003) 643–682 Thauer, R. K., MIBS-39 (2002) 571–619; MILS-2 (2007) 323–356; MILS-6 (2009) 115–132; MILS-6 (2009) 219–240 Thayer, J. S., MIBS-29 (1993) 1–36 Theil, E. C., MIBS-35 (1998) 403–434; MILS-4 (2008) 327–341 Thelander, L., MIBS-30 (1994) 109–129 Thompson, K. H., MIBS-31 (1995) 575–594; MIBS-41 (2004) 221–252 Thorp, H. H., MIBS-33 (1996) 297–324 Thulin, E., MIBS-25 (1989) 255–307 Tilbrook, G. S., MIBS-35 (1998) 691–730 Timmis, K. N., MIBS-28 (1992) 99–156 Tishler, T. A., MILS-1 (2006) 151–177 Tom, R. T., MIBS-23 (1988) 91–121 Toraya, T., MIBS-30 (1994) 217–254 Toth, E., MIBS-40 (2003) 589–641 Tracey, A. S., MIBS-31 (1995) 249–285; MIBS-31 (1995) 287–324 Traynor, D. A., MIBS-37 (2000) 209–278 Treagan, L., MIBS-16 (1983) 27–45 Tsvetkov, Y. D., MIBS-22 (1987) 207–263 Tu, A. T., MIBS-1 (1974) 1–49; MIBS-15 (1983) 193–211 Tuite, E., MIBS-33 (1996) 177–252 Tullius, T. D., MIBS-33 (1996) 453–484 Turner, D. R., MIBS-18 (1984) 137–164 Turnlund, J. R., MIBS-39 (2002) 727–739 Turro, C., MIBS-40 (2003) 323–353

U Udit, A. K., MILS-3 (2007) 157–185 Ukonmaanaho, L., MILS-2 (2007) 1–29 Uldall, A., MIBS-16 (1983) 139–150; MIBS16 (1983) 235–243 Ulstrup, J., MIBS-27 (1991) 57–96 Umezawa, H., MIBS-19 (1985) 81–108 Urade, R., MIBS-38 (2001) 187–196 Urbach, F. L., MIBS-13 (1981) 73–115 Utschig, L. M., MIBS-34 (1997) 503–526

V Vahter, M. E., MIBS-29 (1993) 161–184; MIBS-34 (1997) 371–403 Valensin, D., MILS-1 (2006) 61–87 Valensin, G., MIBS-21 (1987) 1–45; MILS-1 (2006) 61–87 Valenta, J. R., MIBS-19 (1085) 313–333 Valentine, J. S., MIBS-36 (1999) 125–177 Vallee, B. L., MIBS-15 (1983) 1–54; MIBS-25 (1989) 359–394 van der Helm, D., MIBS-35 (1998) 355–401 van der Vlugt, J. I., MILS-2 (2007) 181–239 van Eldik, R., MIBS-32 (1996) 339–378 van Gastel, M., MILS-2 (2007) 279–322 Van Heuvelen, K. M., MILS-6 (2009) 417–460 van Rij, A. M., MIBS-10 (1980) 207–251 van Vliet, A. H. M., MILS-2 (2007) 545–579 Vanquickenborne, L., MIBS-3 (1974) 183–200 Vasˇ a´k, M., MIBS-15 (1983) 213–273; MILS5 (2009) 279–317; MILS-5 (2009) 319–351 Veillon, C., MIBS-16 (1983) 103–122 Veis, A., MILS-4 (2008) 1–35 Vergani, L., MILS-5 (2009) 199–237 Villafranca, J. J., MIBS-4 (1974) 29–59 Vilter, H., MIBS-31 (1995) 325–362 Vol’pin, M. E., MIBS-36 (1999) 485–519 von Ko¨nig, K., MILS-3 (2007) 235–265 Vorholt, J. A., MIBS-39 (2002) 571–619

W Waalkes, M. P., MILS-5 (2009) 399–412 Wackett, L. P., MIBS-28 (1992) 329–356 Wada, O., MIBS-29 (1993) 101–136 Wagner, G. C., MIBS-7 (1978) 241–275 Waite, J. H., MILS-4 (2008) 295–325 Walker, F. A., MIBS-36 (1999) 621–663 Wall, J., MIBS-35 (1998) 667–690 Wallin, S. A., MIBS-27 (1991) 199–222 Wang, J., MIBS-25 (1989) 477–503 Wang, K., MIBS-40 (2003) 707–751 Wang, L., MILS-4 (2008) 413–456 Ward, M. K., MIBS-24 (1988) 217–258 Ward, R. J., MIBS-35 (1998) 633–665; MIBS-41 (2004) 185–219; MILS-1 (2006) 227–279

Met. Ions Life Sci. 2009, 6, 497–510

510 Wasielewski, M. R., MIBS-27 (1991) 361–430 Watanabe, T., MIBS-40 (2003) 161–189 Waterman, M. R., MILS-3 (2007) 361–396 Weatherburn, D. C., MIBS-37 (2000) 209–278 Weatherell, J. A., MIBS-26 (1990) 489–504 Weber, J. H., MIBS-34 (1997) 1–19 Weder, J. E., MIBS-41 (2004) 253–277 Weigand, E., MIBS-15 (1983) 319–361 Weiss, I. M., MILS-4 (2008) 71–126 Welch, R. M., MIBS-26 (1990) 33–56 Welch, T. W., MIBS-33 (1996) 297–324 Wells, G. B., MIBS-22 (1987) 129–206 Weser, U., MIBS-36 (1999) 389–413; MILS-5 (2009) 83–104 Wharton, D. C., MIBS-3 (1974) 157–181 Whitehead, J. P., MIBS-32 (1996) 687–726 Whitehouse, C. J. C., MILS-3 (2007) 437–476 Whitson, L. J., MILS-1 (2006) 179–205 Whittaker, J. W., MIBS-30 (1994) 315–360; MIBS-37 (2000) 587–611 Wilcox, D. E., MIBS-38 (2001) 313–350 Wilkinson, S. R., MIBS-26 (1990) 33–56 Williams, D. R., MIBS-4 (1974) 211–241; MIBS-7 (1978) 29–76; MIBS-12 (1981) 283–317; MIBS-41 (2004) 1–39; MIBS-41 (2004) 139–183 Williams, G., MIBS-21 (1987) 187–227 Williams, J. A. G., MIBS-40 (2003) 233–280 Williams, R. J. P., MIBS-4 (1974) 61–210 Wills, M. R., MIBS-24 (1988) 315–345; MIBS-24 (1988) 347–371 Wilson, I., MIBS-7 (1978) 185–240 Wilson, M. T., MIBS-13 (1981) 187–228 Wilt, F. H., MILS-4 (2008) 37–69 Wink, D. A., MIBS-36 (1999) 547–595 Wink, K., MIBS-41 (2004) 41–69 Winkelmann, G., MIBS-35 (1998) 147–186 Winkler, J. R., MILS-1 (2006) 9–60 Winter, B., MIBS-28 (1992) 157–203 Wireman, J., MIBS-34 (1997) 441–460 Witholt, B., MIBS-28 (1992) 299–327 Witters, R., MIBS-13 (1981) 229–258 Woggon, W.-D., MILS-3 (2007) 27–55

Met. Ions Life Sci. 2009, 6, 497–510

AUTHOR INDEX Wong, H. K. T., MIBS-34 (1997) 131–160 Wong, K. K., MIBS-30 (1994) 279–313 Wong, L. L., MILS-3 (2007) 437–476 Wood, J. M., MIBS-18 (1984) 223–237; MIBS-18 (1984) 333–351 Wu, C.-W., MIBS-15 (1983) 157–192 Wu, F. Y.-H., MIBS-15 (1983) 157–192 Wuthier, R. E., MIBS-17 (1984) 411–472

X Xavier, A. V., MIBS-4 (1974) 61–210; MIBS23 (1988) 285–314 Xia, C., MIBS-42 (2004) 463–497

Y Yamamura, Y., MIBS-29 (1993) 137–160 Yamauchi, O., MIBS-32 (1996) 207–270 Yang, X., MIBS-40 (2003) 707–751 Yano, S., MIBS-32 (1996) 27–60 Yatsimirsky, A. K., MIBS-40 (2003) 369–462 Yoder, D. W., MIBS-37 (2000) 527–557 Yokoyama, A., MIBS-10 (1980) 313–340 Yonetani, T., MIBS-28 (1992) 219–241 Yoshimura, E., MIBS-40 (2003) 161–189 Youdim, M. B. H., MILS-1 (2006) 125–149 Youngs, H. L., MIBS-37 (2000) 559–586 Yuan, J., MIBS-40 (2003) 191–232

Z Zatta, P. F., MILS-1 (2006) 371–393 Zentek, J., MIBS-26 (1990) 57–83 Zerner, B., MIBS-23 (1988) 165–284 Zhang, L., MIBS-41 (2004) 333–378 Zhou, Q., MIBS-41 (2004) 253–277 Zimmermann, W., MIBS-28 (1992) 157–203; MIBS-28 (1992) 357–398 Zot, H. G., MIBS-17 (1984) 381–410 Zuberbu¨hler, A. D., MIBS-5 (1976) 325–368; MIBS-12 (1981) 133–189

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