Nickel and Its Surprising Impact in Nature: Metal Ions in Life Sciences (Vol. 2)

METAL IONS IN LIFE SCIENCES VOLUME 2 Nickel and Its Surprising Impact in Nature METAL IONS IN LIFE SCIENCES edited b...

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METAL IONS IN LIFE SCIENCES VOLUME 2

Nickel and Its Surprising Impact in Nature

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 Zürich Winterthurerstrasse 190 CH-8057 Zürich, Switzerland

VOLUME 2

Nickel and Its Surprising Impact in Nature

Copyright © 2007

John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex PO19 8SQ, England Telephone (+44) 1243 779777 Email (for orders and customer service enquiries): [email protected] Visit our Home Page on www.wileyeurope.com or www.wiley.com All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning or otherwise, except under the terms of the Copyright, Designs and Patents Act 1988 or under the terms of a licence issued by the Copyright Licensing Agency Ltd, 90 Tottenham Court Road, London W1T 4LP, UK, without the permission in writing of the Publisher. Requests to the Publisher should be addressed to the Permissions Department, John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex PO19 8SQ, England, or emailed to [email protected], or faxed to (+44) 1243 770620. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The Publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the Publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. The Publisher, the Editors and the Authors make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of fitness for a particular purpose. This work is sold with the understanding that the Publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for every situation. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of experimental reagents, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each chemical, piece of equipment, reagent, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. The fact that an organization or Website is referred to in this work as a citation and/or a potential source of further information does not mean that the Author, the Editors or the Publisher endorse the information the organization or Website may provide or recommendations it may make. Further, readers should be aware that Internet Websites listed in this work may have changed or disappeared between when this work was written and when it is read. No warranty may be created or extended by any promotional statements for this work. Neither the Publisher nor the Editors nor the Authors shall be liable for any damages arising herefrom. Other Wiley Editorial Offices John Wiley & Sons Inc., 111 River Street, Hoboken, NJ 07030, USA Jossey-Bass, 989 Market Street, San Francisco, CA 94103-1741, USA Wiley-VCH Verlag GmbH, Boschstr. 12, D-69469 Weinheim, Germany John Wiley & Sons Australia Ltd, 42 McDougall Street, Milton, Queensland 4064, Australia John Wiley & Sons (Asia) Pte Ltd, 2 Clementi Loop #02-01, Jin Xing Distripark, Singapore 129809 John Wiley & Sons Canada Ltd, 6045 Freemont Blvd, Mississauga, ONT, L5R 4J3, Canada Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Library of Congress Cataloging-in-Publication Data Nickel and its surprising impact in nature/edited by Astrid Sigel, Helmut Sigel, Roland K. O. Sigel. p. ; cm. – (Metal ions in life sciences ; v. 2) Includes bibliographical references and index. ISBN-13: 978-0-470-01671-8 (cloth : alk. paper) ISBN-10: 0-470-01671-X (cloth : alk. paper) 1. Nickel in the body. 2. Nickel enzymes. 3. Nickel toxicity. I. Sigel, Astrid. II. Sigel, Helmut. III. Sigel, Roland K. O. IV. Series. [DNLM: 1. Nickel–metabolism. 2. Environmental Exposure. 3. Nickel–adverse effects. QU 130 N6323 2006] QP535.N6N53 2006 615.9’25625–dc22 2006022828 British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library ISBN 978-0-470-01671-8 Typeset in 10/12pt Times by Thomson Digital, India Printed and bound in Spain by Grafos S.A. Barcelona This book is printed on acid-free paper responsibly manufactured from sustainable forestry in which at least two trees are planted for each one used for paper production. The figure on the dustcover is Figure 15 of Chapter 4 by Roland K.O. Sigel and Helmut Sigel.

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, such as 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 and we are happy to join forces in this new endeavor with a most experienced publisher in the Sciences, John Wiley & Sons, Ltd, Chichester, UK. 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

PERSPECTIVES OF THE SERIES

(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 on 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 Zürich CH-8057 Zürich Switzerland October 2005

Preface to Volume 2 Nickel and Its Surprising Impact in Nature

This volume is solely devoted to the vibrant research area surrounding nickel and its complexes and their role in Nature. The book opens with the biogeochemistry of this element and its release into the environment, which occurs from both natural and anthropogenic sources, whereby atmospheric distribution plays an important role. In the second chapter the impact of nickel on the metabolism of cyanobacteria and eukaryotic plants including deficiency and toxicity is considered, as is the application of nickel hyperaccumulator plants for phytomining and phytoremediation. Complex formation of nickel(II/III) with amino acids and peptides as well as of nickel(II) with sugar residues, nucleobases, phosphates, nucleosides, and nucleic acids is summarized in Chapters 3 and 4, respectively, by also taking into account intramolecular equilibria and comparisons with related metal ions. Bioinspired nickel coordination chemistry has flourished in recent years and the resulting synthetic models for the active sites of nickel-containing enzymes are reviewed in Chapter 5. In fact, each of the well-established biological nickel sites is rather unique with respect to its structure and function. Hence, the following eight chapters are individually devoted to the various nickel enzymes which catalyze rather diverse reactions. For example, urease reduces the half life of urea in water from about 3.6 years to a few microseconds, whereas nickeliron hydrogenases catalyze the heterolytic conversion of dihydrogen into protons and electrons and vice versa. Next, methyl-coenzyme M reductase and its nickel corphin coenzyme F430 in methanogenic archaea are described in detail as are acetyl-coenzyme A synthases and nickel-containing carbon monoxide dehydrogenases. These critical reviews are followed by in-depth considerations on nickel superoxide dismutase, and the nickel-dependent glyoxalase I enzymes. The role of nickel in acireductone dioxygenase and the properties of the nickel-regulated peptidyl-prolyl cis/trans isomerase SlyD are discussed next.

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PREFACE TO VOLUME 2

Nickel is toxic to cells and therefore the synthesis of nickel enzymes requires carefully controlled nickel-processing mechanisms that range from selective transport of nickel into the cells to productive insertion of nickel into the correct apoproteins. This demanding task is the focus of Chapter 14 which is devoted to the chaperones of nickel metabolism. The primary colonization and long-term survival of Helicobacter pylori in the hostile gastric niche and the role of nickel in this environmental adaptation is covered in detail in Chapter 15. Nickel is widely employed in modern industry in conjunction with other metals for the production of alloys for coins, jewellery, and stainless steel; it is also used for plating, battery production, as a catalyst, etc. Workers are exposed to nickel at all stages of the processing of nickel-containing products through air, water or skin contacts. For example, the exposure to airborne nickel-containing particles has long been known to cause acute respiratory symptoms ranging from mild irritation and inflammation of the respiratory system to bronchitis, asthma, and pulmonary fibrosis and edema. Another well-known adverse effect is allergic contact dermatitis. The indicated health problems caused by nickel exposure are mediated by an active change in the expression of genes that control inflammation, the response to stress, cell proliferation or cell death. All this and more is covered in Chapter 16. However, the most serious health effects beyond nickel toxicity relate to carcinogenesis; these concerns represent an area of considerable research activity today as is evident from the terminating chapter of Nickel and Its Surprising Impact in Nature. Astrid Sigel Helmut Sigel Roland K. O. Sigel

Contents

HISTORICAL DEVELOPMENT AND PERSPECTIVES OF THE SERIES PREFACE TO VOLUME 2 CONTRIBUTORS TO VOLUME 2 TITLES OF VOLUMES 1–44 IN THE METAL IONS IN BIOLOGICAL SYSTEMS SERIES CONTENTS OF VOLUMES IN THE METAL IONS IN LIFE SCIENCES SERIES

1 BIOGEOCHEMISTRY OF NICKEL AND ITS RELEASE INTO THE ENVIRONMENT

v vii xvii xxi xxiii

1

Tiina M. Nieminen, Liisa Ukonmaanaho, Nicole Rausch, and William Shotyk 1. 2. 3. 4. 5. 6. 7. 8.

Introduction Chemistry of Nickel Ancient and Modern Uses of Nickel Sources of Atmospheric Nickel Deposition and Fate of Atmospheric Nickel Historical Records of Nickel Deposition Bioavailability and Mobility of Nickel in Soils Summary and Conclusions Abbreviations References

2 2 6 6 7 14 17 21 21 22

x

CONTENTS

2 NICKEL IN THE ENVIRONMENT AND ITS ROLE IN THE METABOLISM OF PLANTS AND CYANOBACTERIA

31

Hendrik Küpper and Peter M. H. Kroneck 1. 2. 3. 4. 5.

Introduction Nickel as a Micronutrient for Plants and Cyanobacteria Nickel as an Environmental Pollutant and Its Effects on Plants Nickel Hyperaccumulation Outlook Acknowledgments Abbreviations References

3 NICKEL ION COMPLEXES OF AMINO ACIDS AND PEPTIDES

32 37 40 48 53 54 54 55

63

Teresa Kowalik-Jankowska, Henryk Kozlowski, Etelka Farkas, and Imre Sóvágó 1. Introduction 2. Complexes of Amino Acids and Derivatives 3. Complexes of Peptides and Related Ligands 4. Formation of Nickel(II) Complexes under Biological Conditions: Model Calculations in Multicomponent Systems 5. Conclusions Abbreviations References

4 COMPLEX FORMATION OF NICKEL(II) AND RELATED METAL IONS WITH SUGAR RESIDUES, NUCLEOBASES, PHOSPHATES, NUCLEOTIDES, AND NUCLEIC ACIDS

64 66 76 94 96 97 98

109

Roland K. O. Sigel and Helmut Sigel 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Introduction Nickel(II)–Sugar Interactions Interactions of Nickel(II) with Nucleobase Residues Complexes of Nickel(II) with Phosphates Nickel(II) Complexes of Nucleotides Complexes of Some Less Common Nucleotides Complexes of Some Nucleotide Derivatives and Analogs Mixed Ligand Complexes Containing a Nucleotide Nickel(II) Binding in Nucleic Acids Concluding Remarks

110 112 118 128 131 144 149 159 165 168

CONTENTS

Acknowledgments Abbreviations and Definitions References

5 SYNTHETIC MODELS FOR THE ACTIVE SITES OF NICKEL-CONTAINING ENZYMES

xi

169 169 172

181

Jarl Ivar van der Vlugt and Franc Meyer 1. 2. 3. 4. 5. 6.

Introduction Models for Cofactor F430 Models for Sulfur-Rich Nickel Sites Models for the Urease Active Site Models for Acireductone Reductase Concluding Remarks Acknowledgments Abbreviations References

6 UREASE: RECENT INSIGHTS ON THE ROLE OF NICKEL

181 182 191 214 229 230 230 230 232

241

Stefano Ciurli 1. 2. 3. 4. 5.

Introduction: Urease and Its Biological Significance The Biochemistry of Urease Structural Studies on Bacterial Ureases The Structure-Based Mechanism of Urease Conclusions Acknowledgments Abbreviations References

7 NICKEL IRON HYDROGENASES

242 243 244 267 273 273 273 274

279

Wolfgang Lubitz, Maurice van Gastel, and Wolfgang Gärtner 1. 2. 3. 4. 5. 6. 7.

Introduction to Hydrogenases Biochemistry and Molecular Biology Crystallization and X-Ray Structure Analysis Spectroscopic Investigations Electrochemistry Hydrogenase Function and the Catalytic Cycle Conclusions and Outlook Acknowledgments Abbreviations References

280 283 291 295 306 309 312 314 314 315

xii

CONTENTS

8 METHYL-COENZYME M REDUCTASE AND ITS NICKEL CORPHIN COENZYME F430 IN METHANOGENIC ARCHAEA

323

Bernhard Jaun and Rudolf K. Thauer 1. Introduction 2. Structure and Properties of Coenzyme F430 3. Molecular Properties of Methyl-Coenzyme M Reductase 4. Catalytic Properties of Methyl-Coenzyme M Reductase Acknowledgments Abbreviations References

324 328 335 345 350 350 351

9 ACETYL-COENZYME A SYNTHASES AND NICKEL-CONTAINING CARBON MONOXIDE DEHYDROGENASES

357

Paul A. Lindahl and David E. Graham 1. Introduction 2. Structure and Function of Carbon Monoxide Dehydrogenases 3. Sequence Analysis and Phylogeny of Carbon Monoxide Dehydrogenases 4. Acetyl-Coenzyme A Synthases/Carbon Monoxide Dehydrogenases 5. Sequence Analysis and Phylogeny of the α Subunit 6. Corrinoid Iron-Sulfur Proteins 7. Acetyl-Coenzyme A Decarbonylase/Synthases 8. Physiological Roles and Evolution of Acetyl-Coenzyme A Synthase/Carbon Monoxide Dehydrogenase Proteins 9. Origins and Evolution of ACDS, ACS/CODH, and CODH Complexes Acknowledgments Abbreviations Appendices References

10 NICKEL SUPEROXIDE DISMUTASE

358 361 369 373 381 382 384 386 392 393 394 395 411

417

Peter A. Bryngelson and Michael J. Maroney 1. Introduction 2. Molecular Biology 3. Structural Biology 4. Mechanism

418 422 426 429

CONTENTS

5.

Conclusions Acknowledgments Abbreviations and Definitions References

11 BIOCHEMISTRY OF THE NICKEL-DEPENDENT GLYOXALASE I ENZYMES

xiii

438 438 439 439

445

Nicole Sukdeo, Elisabeth Daub, and John F. Honek 1. 2. 3. 4. 5.

Introduction Biochemical Investigations of Glyoxalase I Biophysical and Mechanistic Studies of Glyoxalase I Glyoxalase I Genes and Protein Sequence Comparisons Glyoxalase I as a Member of the βαβββ Superfamily of Proteins 6. Other Aspects of Glyoxalase I 7. Conclusions Acknowledgments Abbreviations References

12 NICKEL IN ACIREDUCTONE DIOXYGENASE

446 450 453 460 463 464 465 466 466 467

473

Thomas C. Pochapsky, Tingting Ju, Marina Dang, Rachel Beaulieu, Gina M. Pagani, and Bo OuYang 1. Introduction 2. The Methionine Salvage Pathway 3. One Protein, Two Enzymes: Acireductone Dioxygenase from Klebsiella oxytoca 4. Homologs of Acireductone Dioxygenase from Other Organisms 5. Known Acireductone Dioxygenase Structures 6. Spectroscopic Probes of Acireductone Dioxygenase Enzyme Active Sites 7. Enzymatic Studies of Acireductone Dioxygenase 8. Mechanistic Considerations: What is the Role of Ni(II) in Acireductone Dioxygenase Activity? 9. Structurally and Functionally Related Enzymes 10. Future Directions Acknowledgments Abbreviations References

474 475 477 481 483 486 489 490 493 495 497 497 498

xiv

CONTENTS

13 THE NICKEL-REGULATED PEPTIDYL PROLYL CIS/TRANS ISOMERASE SlyD

501

Frank Erdmann and Gunter Fischer 1. Introduction 2. SlyD Belongs to the Peptidyl Prolyl cis/trans Isomerases 3. Insights into the Biological Role of SlyD 4. Conclusions Abbreviations and Definitions References

14 CHAPERONES OF NICKEL METABOLISM

502 503 513 515 515 516

519

Soledad Quiroz, Jong K. Kim, Scott B. Mulrooney, and Robert P. Hausinger 1. Introduction to Nickel Metabolism 2. Nickel Metallochaperones 3. Molecular Chaperones Involved in Nickel Metabolism 4. Conclusions and Remaining Questions Acknowledgments Abbreviations References

15 THE ROLE OF NICKEL IN ENVIRONMENTAL ADAPTATION OF THE GASTRIC PATHOGEN HELICOBACTER PYLORI

520 530 534 538 539 539 540

545

Florian D. Ernst, Arnoud H. M. van Vliet, Manfred Kist, Johannes G. Kusters, and Stefan Bereswill 1. 2. 3. 4. 5. 6. 7.

Introduction Nickel Enzymes and Environmental Adaptation Nickel Uptake Systems Mechanisms of Nickel Regulation Protection of Nickel Metabolism Metal Metabolism as Drug Target: Therapeutic Considerations Conclusions Abbreviations References

16 NICKEL-DEPENDENT GENE EXPRESSION

546 554 560 562 566 568 570 570 571

581

Konstantin Salnikow and Kazimierz S. Kasprzak 1. Introduction 2. Genetic and Epigenetic Changes in Nickel-Exposed Cells

582 585

CONTENTS

3. 4. 5. 6. 7. 8.

Alteration of Gene Expression Following Nickel-Induced Lung Injury Nickel-Induced Allergy and Gene Expression Nickel-Induced Expression of Erythropoietin Alteration of Transcription Factors and Signaling Pathways Changes in Gene Expression and Nickel Carcinogenesis Conclusions Acknowledgments Abbreviations References

17 NICKEL TOXICITY AND CARCINOGENESIS

xv

587 592 592 593 607 608 609 609 610

619

Kazimierz S. Kasprzak and Konstantin Salnikow 1. Introduction 2. An Overview of Nickel Toxicity 3. Nickel-Induced Carcinogenesis 4. Conclusion Acknowledgments Abbreviations References SUBJECT INDEX

621 621 638 645 647 647 647 661

Contributors

Numbers in parentheses indicate the pages on which the authors’ contributions begin. Rachel Beaulieu Departments of Chemistry and Biochemistry, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454-9110, USA (473) Stefan Bereswill Charité - University Medicine Berlin, Institute for Microbiology and Hygiene, Campus Charité Mitte, Dorotheenstrasse 96, D-10117 Berlin, Germany, ⬍[email protected]⬎ (545) Peter A. Bryngelson Department of Chemistry, 701 Lederle Graduate Research Tower, University of Massachusetts, 710 North Pleasant Street, Amherst, MA 01003-9336, USA, ⬍[email protected]⬎ (417) Stefano Ciurli Laboratory of Bioinorganic Chemistry, Department of AgroEnvironmental Science and Technology, University of Bologna, Viale Giuseppe Fanin 40, I-40127 Bologna, Italy, ⬍[email protected]⬎ (241) Marina Dang Departments of Chemistry and Biochemistry, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454-9110, USA (473) Elisabeth Daub Department of Chemistry, University of Waterloo, Waterloo, Ontario, N2L 3G1, Canada, ⬍[email protected]⬎ (445) Frank Erdmann Max-Planck Research Unit for Enzymology of Protein Folding, Weinbergweg 22, D-06120 Halle, Germany, ⬍[email protected]⬎ (501) Florian D. Ernst Department of Gastroenterology and Hepatology, Erasmus MC - University Medical Center, Rotterdam, The Netherlands (545)

xviii

CONTRIBUTORS

Etelka Farkas Inorganic and Analytical Chemistry Department, University of Debrecen, Egyetem ter. 1, P.O. Box 21, H-4010 Debrecen, Hungary, ⬍[email protected]⬎ (63) Gunter Fischer Max-Planck Research Unit for Enzymology of Protein Folding, Weinbergweg 22, D-06120 Halle, Germany, ⬍[email protected]⬎ (501) Wolfgang Gärtner Max-Planck-Institute for Bioinorganic Chemistry, Stiftstrasse 34-36, D-45470 Mülheim/Ruhr, Germany (279) David E. Graham Department of Chemistry and Biochemistry, University of Texas, Austin, TX 78712, USA, ⬍[email protected]⬎ (357) Robert P. Hausinger Department of Biochemistry and Molecular Biology, Cell and Molecular Biology Program, and Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI 48824-4320, USA, ⬍[email protected]⬎ (519) John F. Honek Department of Chemistry, University of Waterloo, Waterloo, Ontario, N2L 3G1, Canada, ⬍[email protected]⬎ (445) Bernhard Jaun Organic Chemistry, ETHZ, ETH Hönggerberg HCI E317, CH-8093 Zürich, Switzerland, ⬍[email protected]⬎ (323) Tingting Ju Departments of Chemistry and Biochemistry, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454-9110, USA (473) Kazimierz S. Kasprzak Laboratory for Comparative Carcinogenesis, National Cancer Institute at Frederick, Bldg 538, Room 205E, Frederick, MD 21702-1201, USA, ⬍[email protected]⬎ (582, 619) Jong K. Kim Cell and Molecular Biology Program, Michigan State University, East Lansing, MI 48824-4320, USA (519) Manfred Kist Department of Microbiology and Hygiene, Institute of Medical Microbiology and Hygiene, University Hospital, Freiburg, Germany (545) Teresa Kowalik-Jankowska Faculty of Chemistry, University of Wroclaw, F. JoliotCurie 14, PL-50383 Wroclaw, Poland, ⬍[email protected]⬎ (63) Henryk Kozlowski Faculty of Chemistry, University of Wroclaw, F. JoliotCurie 14, PL-50383 Wroclaw, Poland, ⬍[email protected]⬎ (63)

CONTRIBUTORS

xix

Peter M. H. Kroneck Fachbereich Biologie, Universität Konstanz, Postfach M665, D-78457 Konstanz, Germany, ⬍[email protected]⬎ (31) Hendrik Küpper Fachbereich Biologie, Universität Konstanz, Postfach M665, D-78457 Konstanz, Germany, ⬍[email protected]⬎ (31) Johannes G. Kusters Department of Gastroenterology and Hepatology, Erasmus MC-University Medical Center, Rotterdam, The Netherlands (545) Paul A. Lindahl Department of Chemistry and of Biochemistry and Biophysics, Texas A&M University, College Station, TX 77843-3255, USA, ⬍[email protected]⬎ (357) Wolfgang Lubitz Max-Planck-Institute for Bioinorganic Chemistry, Stiftstrasse 34-36, D-45470 Mülheim/Ruhr, Germany, ⬍[email protected]⬎ (279) Michael J. Maroney Department of Chemistry, 701 Lederle Graduate Research Tower, University of Massachusetts, 710 North Pleasant Street, Amherst, MA 01003-9336, USA, ⬍[email protected]⬎ (417) Franc Meyer Institut für Anorganische Chemie, Georg-August-Universität Göttingen, Tammannstrasse 4, D-37077 Göttingen, Germany, ⬍[email protected]⬎ (182) Scott B. Mulrooney Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI 48824-4320, USA (519) Tiina M. Nieminen Finnish Forest Research Institute (Metla), P. O. Box 18, FI-01301 Vantaa, Finland, ⬍[email protected]⬎ (1) Bo OuYang Departments of Chemistry and Biochemistry, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454-9110, USA (473) Gina M. Pagani Departments of Chemistry and Biochemistry, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454-9110, USA (473) Thomas C. Pochapsky Departments of Chemistry and Biochemistry, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454-9110, USA, ⬍[email protected]⬎ (473) Soledad Quiroz Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824-4320, USA (519)

xx

CONTRIBUTORS

Nicole Rausch Institute of Environmental Geochemistry, University of Heidelberg, Im Neuenheimer Feld 236, D-69120 Heidelberg, Germany, ⬍[email protected]⬎ (1) Konstantin Salnikow Laboratory for Comparative Carcinogenesis, National Cancer Institute at Frederick, Bldg 538, Room 205E, Frederick, MD 21702-1201, USA, ⬍[email protected]⬎ (582, 619) William Shotyk Institute of Environmental Geochemistry, University of Heidelberg, Im Neuenheimer Feld 236, D-69120 Heidelberg, Germany, ⬍[email protected]⬎ (1) Helmut Sigel Department of Chemistry, Inorganic Chemistry, University of Basel, Spitalstrasse 51, CH-4056 Basel, Switzerland, ⬍[email protected]⬎ (109) Roland K. O. Sigel Institute of Inorganic Chemistry, University of Zürich, Room 34-F-36, Winterthurerstrasse 190, CH-8057 Zürich, Switzerland, ⬍[email protected]⬎ (109) Imre Sóvágó Inorganic and Analytical Chemistry Department, University of Debrecen, Egyetem ter. 1, P.O. Box 21, H-4010 Debrecen, Hungary, ⬍[email protected]⬎ (63) Nicole Sukdeo Department of Chemistry, University of Waterloo, Waterloo, Ontario, N2L 3G1, Canada, ⬍[email protected]⬎ (445) Rudolf K. Thauer Max Planck Institute for Terrestrial Microbiology, Karlvon-Frisch-Strasse, D-35043 Marburg, Germany, ⬍[email protected]⬎ (323) Liisa Ukonmaanaho Finnish Forest Research Institute (Metla), P. O. Box 18, FI-01301 Vantaa, Finland, ⬍[email protected]⬎ (1) Jarl Ivar van der Vlugt Institut für Anorganische Chemie, Georg-AugustUniversität Göttingen, Tammannstrasse 4, D-37077 Göttingen, Germany, ⬍[email protected]⬎ (182) Maurice van Gastel Max-Planck-Institute for Bioinorganic Chemistry, Stiftstrasse 34-36, D-45470 Mülheim/Ruhr, Germany (279) Arnoud H. M. van Vliet Department of Gastroenterology and Hepatology, Erasmus MC - University Medical Center, Rotterdam, The Netherlands (545)

Titles of Volumes 1–44 in the Metal Ions in Biological Systems Series edited by the SIGELs and published by Dekker/Taylor & Francis 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 17: Volume 18: Volume 19: Volume 20: Volume 21: Volume 22: Volume 23: Volume 24: Volume 25:

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 Aluminum and Its Role in Biology Interrelations Among Metal Ions, Enzymes, and Gene Expression

xxii

VOLUMES IN THE MIBS SERIES

Volume 26: Compendium on Magnesium and its Role in Biology, Nutrition, and Physiology Volume 27: Electron Transfer Reactions in Metalloproteins Volume 28: Degradation of Environmental Pollutants by Microorganisms and Their Metalloenzymes Volume 29: Biological Properties of Metal Alkyl Derivatives Volume 30: Metalloenzymes Involving Amino Acid-Residue and Related Radicals Volume 31: Vanadium and Its Role for Life Volume 32: Interactions of Metal Ions with Nucleotides, Nucleic Acids, and Their Constituents Volume 33: Probing Nucleic Acids by Metal Ion Complexes of Small Molecules Volume 34: Mercury and Its Effects on Environment and Biology Volume 35: Iron Transport and Storage in Microorganisms, Plants, and Animals Volume 36: Interrelations Between Free Radicals and Metal Ions in Life Processes Volume 37: Manganese and Its Role in Biological Processes Volume 38: Probing of Proteins by Metal Ions and Their Low-Molecular-Weight Complexes Volume 39: Molybdenum and Tungsten. Their Roles in Biological Processes Volume 40: The Lanthanides and Their Interrelations with Biosystems Volume 41: Metal Ions and Their Complexes in Medication Volume 42: Metal Complexes in Tumor Diagnosis and as Anticancer Agents Volume 43: Biogeochemical Cycles of Elements Volume 44: Biogeochemistry, Availability, and Transport of Metals in the Environment

Contents of Volumes in the Metal Ions in Life Sciences Series edited by the SIGELs and published by John Wiley & Sons, Ltd, Chichester, UK Volume 1: Neurodegenerative Diseases and Metal Ions 1. The Role of Metal Ions in Neurology. An Introduction Dorothea Strozyk and Ashley I. Bush 2. Protein Folding, Misfolding, and Disease Jennifer C. Lee, Judy E. Kim, Ekaterina V. Pletneva, Jasmin Faraone-Mennella, Harry B. Gray, and Jay R. Winkler 3. Metal Ion Binding Properties of Proteins Related to Neurodegeneration Henryk Kozlowski, Marek Luczkowski, Daniela Valensin, and Gianni Valensin 4. Metallic Prions: Mining the Core of Transmissible Spongiform Encephalopathies David R. Brown 5. The Role of Metal Ions in the Amyloid Precursor Protein and in Alzheimer’s Disease Thomas A. Bayer and Gerd Multhaup 6. The Role of Iron in the Pathogenesis of Parkinson’s Disease Manfred Gerlach, Kay L. Double, Mario E. Götz, Moussa B. H. Youdim, and Peter Riederer 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

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CONTENTS OF MILS VOLUMES

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 Tamás 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:

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

8. 9.

10.

Nickel and Its Surprising Impact in Nature this book

The Ubiquitous Roles of Cytochrome P450 Proteins (in press) Diversities and Similarities of P450 Systems: An Introduction Mary A. Schuler and Stephen G. Sligar Structural and Functional Mimics of Cytochromes P450 Wolf-D. Woggon Structures of P450 Proteins and Their Molecular Phylogeny Thomas L. Poulos and Yergalem T. Meharenna Aquatic P450 Species Mark J. Snyder The Electrochemistry of Cytochrome P450 Systems 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 König 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


xxv

11. Cytochrome P450-Catalyzed Hydroxylations and Epoxidations Roshan Perera, Shengxi Jin, Masanori Sono, and John H. Dawson 12. Cytochrome P450 and Steroid Hormone Biosynthesis Rita Bernhardt and Michael R. Waterman 13. Carbon-Carbon Bond Cleavage by P450 Systems James J. De Voss and Max J. Cryle 14. Design and Engineering of Cytochrome P450 Systems Stephen G. Bell, Nicola Hoskins, Christopher J. C. Whitehouse, and Luet L. Wong 15. Chemical Defence and Exploitation: Biotransformation of Xenobiotics by Cytochrome P450 Enzymes Elizabeth M. J. Gillam and Dominic J. B. Hunter 16. Drug Metabolism as Catalyzed by Human Cytochrome P450 Systems F. Peter Guengerich 17. Cytochrome P450 Enzymes: Observations from the Clinic Peggy L. Carver Subject Index

Volume 4: 1. 2.

3. 4. 5. 6. 7. 8.

9.

Biomineralization. From Nature to Application (tentative contents) Crystals and Life. An Introduction Arthur Veis Gene-Directed Crystal Growth Exemplified by the Biomineralization of Calcium Carbonate Fred H. Wilt and Christopher E. Killian The Role of Enzymes in Biomineralization Processes Ingrid M. Weiss and Frédéric Marin Metal-Bacterial Interactions at Both the Planktonic Cell and Biofilm Levels Ryan Hunter and Terry J. Beveridge Biomineralization of Calcium Carbonate. The Interplay with Biosubstrates Amir Berman and Yael Levi-Kalisman Sulfate-Containing Biominerals Fabienne Bosselmann and Matthias Epple Oxalate Biominerals Enrique J. Baran and Paula V. Monje Structural Control, Molecular Components, and Multi-Level Regulation of Biosilification in Diatoms Aubrey K. Davis, Kim Thamatrakoln, and Mark Hildebrand Dynamics of Biomineralization and Biodemineralization Lijun Wang and George H. Nancollas

xxvi


10. Mechanism of Mineralization of Collagen Based Connective Tissue Adele J. Boskey 11. Mammalian Enamel Formation Janet Moradian-Oldak and Michael Paine 12. Heavy Metals in the Jaws of Invertebrates Helga C. Lichtenengger 13. Ferritin. Biomineralization of Iron Elizabeth C. Theil 14. Molecular Biology and Magnetism for Magnetic Iron Minerals in Bacteria Richard B. Frankel, Sabrina Schübbe, and Dennis Bazylinski 15. Mechanical Design of Biomineralized Tissues Peter Fratzl 16. Biominerals. Recorders of the Past Danielle Fortin and Susan Glasauer 17. Bio-Inspired Growth of Mineralized Tissue Darilis Suarez and William L. Murphy 18. Biomineralization of Novel Inorganic Materials for Application Helmut Cölfen and Markus Antonietti 19. Crystal Tectonics. Chemical Construction and Self-Organization Annie K. Powell Subject Index Comments and suggestions with regard to contents, topics, and such for future volumes of the series are welcome.

Met. Ions Life Sci. 2, 1–30 (2007)

1 Biogeochemistry of Nickel and Its Release into the Environment Tiina M. Nieminen,1 Liisa Ukonmaanaho,1 Nicole Rausch, 2 and William Shotyk2 1

2

Finnish Forest Research Institute (Metla), FI-01301 Vantaa, Finland

Institute of Environmental Geochemistry, University of Heidelberg, D-69120 Heidelberg, Germany <[email protected]>

1. INTRODUCTION 2. CHEMISTRY OF NICKEL 2.1. Chemical Properties 2.2. Geological Abundance and Occurrence 2.3. Measurement of Nickel in Environmental Samples 3. ANCIENT AND MODERN USES OF NICKEL 4. SOURCES OF ATMOSPHERIC NICKEL 4.1. Natural Sources 4.2. Anthropogenic Sources 5. DEPOSITION AND FATE OF ATMOSPHERIC NICKEL 5.1. Air Quality and Deposition in Polluted Areas versus Remote Areas 5.2. Regional Indicator Surveys: The Use of Mosses, Lichens, Bark 5.2.1. Mosses 5.2.2. Lichens 5.2.3. Bark Metal Ions in Life Sciences, Volume 2 © 2007 John Wiley & Sons, Ltd

Edited by Astrid Sigel, Helmut Sigel and Roland K. O. Sigel

2 2 2 4 4 6 6 6 7 7 7 8 8 9 10

2

NIEMINEN, SHOTYK et al.

5.3. Nickel Concentration in Aquatic and Biotic Media 5.4. Nickel Fluxes in Forested Catchments 6. HISTORICAL RECORDS OF NICKEL DEPOSITION 6.1. Sediment 6.2. Peat Bog 6.3. Polar Snow and Ice 7. BIOAVAILABILITY AND MOBILITY OF NICKEL IN SOILS 7.1. Uptake and Translocation of Nickel by Plants 7.2. Importance of Partitioning for Bioavailability and Mobility 8. SUMMARY AND CONCLUSIONS ABBREVIATIONS REFERENCES

1.

10 12 14 14 15 15 17 17 19 21 21 22

INTRODUCTION

Nickel and its compounds are released into the atmosphere from both natural and anthropogenic sources. Although Ni is an essential element to plants and many other biota, there has been much more concern about the toxicity of Ni than about Ni deficiency. Field observations have indicated a significant increase in heavy metal concentrations in agricultural and forest soils as well as in marine and inland water sediments during the last century [1,2]. An increase is also frequently observed in remote areas thousands of kilometers away from major anthropogenic sources due to long-range atmospheric transport, e.g., elevated Ni concentrations have been reported from the Norwegian arctic and the Finnish Lapland region [3–5]. Nriagu and Pacyna [6] give an estimation of 24 000–87 000 tons in 1983 for the worldwide Ni emissions to the atmosphere, and for the total global release of Ni into soils (atmospheric fallout, wastes, fertilizers, sewage sludge, etc.) an estimate ranging from 106 000 to 544 000 tons per year. In this context, it is important to know the background concentrations of trace elements in uncontaminated sites for comparison with polluted areas.

2. 2.1.

CHEMISTRY OF NICKEL Chemical Properties

Nickel (Z 28, atomic weight 58.69) belongs to Group 10 (formerly VIII) of the periodic table, the so-called iron–cobalt–nickel group of metals. As such, Met. Ions Life Sci. 2, 1–30 (2007)

Ni IN BIOGEOCHEMISTRY AND ENVIRONMENT

3

1.2 System Ni – O – H – S 25°C, 1 bar PO = 2 1b ar

1.0 0.8 0.6

Ni(OH)2

Eh (V)

0.2 0.0 –0.2

HNiO2–

Ni2+

0.4

NiS

PH

–0.4

2

=1

bar

–0.6 –0.8 0

2

4

6

8

10

12

14

pH

Figure 1. Eh–pH diagram for part of the system Ni–O–H–S. Assumed activities for dissolved species are [Ni2] 106, [S2–] 103. Redrawn from [9] with kind permission of Springer Science and Business Media.

Ni is closely related to Co in both its chemical and biochemical properties. The electronic configuration is (Ar) 3d8 4s2. Normally, Ni occurs either in the 0 or II oxidation states, although the I and III states can exist under certain conditions. As the latter ions are not stable in aqueous solution [7], redox processes are not directly important in the environment. Elemental Ni is a silvery white, hard, malleable metal which melts at 1453C and has a density of 8.9 g cm1. Five stable isotopes exist of Ni (with natural abundances): 58Ni (67.9%), 60Ni (26.2%), 61 Ni (1.19%), 62Ni (3.66%), and 64Ni (1.08%). There are 18 radioisotopes, with the most stable being 59Ni (half-life 76 000 yr) and 63Ni (half-life 100 yr). Nickel readily loses two electrons, yielding Ni2. This is the dominant inorganic species throughout the pH and Eh range of most natural waters (Figure 1). However, in the presence of dissolved organic matter, Ni can form strong complexes with organic ligands. In increasingly calcareous/alkaline waters it is stable as NiHCO3 and NiCO30, and Ni(OH)20, respectively. In zones of active bacterial sulfate reduction such as anoxic sediments, the solubility of Ni is regulated by the formation of Ni sulfide (NiS). In the presence of phosphate, as in fertilized agricultural soils, Ni3 (PO4)2 can be formed resulting in very low Ni2 concentrations in the aqueous phase (solubility product about 1032). In most soils, Ni is bound to ion Met. Ions Life Sci. 2, 1–30 (2007)

4


exchange sites, is specifically adsorbed, or adsorbed on or coprecipitated with aluminum and iron oxyhydroxides. These are the dominant processes in neutral to alkaline soils. In acidic organic-rich soils, where fulvic and humic acids are formed by the decomposition of organic material, Ni may be quite mobile, possibly because of complexation by these ligands [8].

2.2.

Geological Abundance and Occurrence

The abundance of Ni in the earth’s crust is ⬃80 µg g1 [7]. While concentrations in most rocks are below 150 µg g1 (sedimentary rocks: 70 µg g1; basalts: 150 µg g1; granitic rocks: 15 µg g1), Ni is highly enriched in ultramafic rocks such as peridotite and serpentine (2000 µg g1) [10], as well as meteorites (5–20% Ni). Nickel as a chalcophilic element forms many different sulfides, arsenides as well as antimonides in nature (e.g., NiS, NiAs, NiAs2–3, NiAsS, NiSb, NiSbS). There exist two different types of commercially exploitable deposits: sulfide ores, with pyrrhotite, pentlandite, pyrite and chalcopyrite as the main Ni-containing minerals, and laterites, formed during tropic weathering, such as garnierite ((Mg,Ni)3 (OH) 4 [Si2O5]). The latter can often be mined in open pits. The main Ni producers are Canada (sulfidic ores at the Sudbury and Thompson mines), Russia (laterites and pentlandite at Norilsk and Pechenga), and Indonesia and Australia (laterites).

2.3. Measurement of Nickel in Environmental Samples The concentrations of total Ni in uncontaminated environmental samples range from 0.6 pg g1 in ice-core samples [11] to more than 1 mg g1 in serpentine soils [7]. The most common techniques for the quantification of Ni in these samples are atomic absorption spectrometry (AAS), inductively coupled plasma optical emission spectrometry (ICP-OES), and inductively coupled plasma-mass spectrometry (ICP-MS). Approved test methods are given by the Environmental Protection Agency (EPA) [12–14]. Limits of detection (LODs) of 40 µg L1 are reported for the analysis of water samples using flame AAS, and 1 µg L1 using graphite furnace AAS [12,13]. A great advantage of ICP-OES and ICP-MS is their capability of multi-element analysis. An additional advantage of ICP-MS is the low LOD for most elements (lower µg L1 range). Although these methods are suitable for acid digestions of soil and sediment samples as well as many water samples, the Ni concentration in some natural and drinking waters is often below the LODs of the ICP-MS. Therefore, the EPA specifically developed three different preconcentration protocols for the determination of metals in ambient waters at EPA water quality criteria levels (WQC), resulting in a LOD of 0.029 µg L1 for Ni [14]. Met. Ions Life Sci. 2, 1–30 (2007)


5

However, one of the greatest difficulties in measuring Ni at these levels is precluding sample contamination during collection, transport, and analysis. Over the past decades, marine chemists have come to recognize that much of the historical data on the concentrations of Ni in seawater are erroneously high because the concentrations reflect contamination from sampling and analysis rather than ambient levels [14]. Therefore, the EPA developed a clean sampling procedure of ambient water [14]. Solid phase extraction is a very sensitive, fast and economic preconcentration method [15], and various extractants are reported in the recent literature [15–24]. Alternatively, Ni can be preconcentrated using coprecipitation [25,26]. With the development of high-resolution ICP-MS instruments and the installation of clean lab facilities, a new chapter in Ni quantification has been opened. The main advantages compared with quadrupole ICP-MS instruments are: (1) increased sensitivity (factor 10); (2) low background intensities; (3) the separation of Ni from polyatomic interferences (CaO, CaOH); and (4) the small sample volumes required. This allowed the multi-element analysis of bog porewater [27] where preconcentration procedures were not possible due to the low sample amounts available. Due to the low background levels of the instrument, LODs are dependent on the effort to avoid random sample contamination. Using class 100 flow benches and high-purity materials (Teflon, Peek), e.g., LODs of 15 ng L1 could be obtained for Ni [27]. Using the slurry-sample introduction technique in AAS [28,29] and laserablation ICP-MS [29], the direct determination of Ni in solid environmental samples, without time-consuming digestion procedures, is possible. The LODs attainable with laser-ablation ICP-MS depend on the size of the ablation pits, but are typically in the low to sub ng g1 range for pits of 100 µm [30]. However, XRF spectrometry is a cheap, fast and simple alternative for the multi-element analysis of solid samples at the µg g1 concentration level, and it is non-destructive. Using total reflexion XRF, aqueous solutions can be analyzed [31,32]. For analyzing small samples, such as single mineral grains, electron microprobe analyzers (EMPA) have been used extensively, although LODs are in general high (50–200 µg g1 for many elements) [33]. Synchrotron-based XRF allows the establishment of elemental mappings of small-scale samples, with lateral resolutions in the µm range [34]. By combining different synchrotron X-ray techniques, such as X-ray diffraction, X-ray absorption fine-structure analysis (XAFS), and X-ray absorption near edge structure analysis (XANES), information on the mineral phase, the oxidation state and the nearest neighbour atom can be obtained [35–37]. For the speciation of Ni in aqueous solutions, adsorptive cathodic stripping voltammetry is frequently employed [38–40]. Using chromatographic methods both speciation and the separation of different metals in aqueous solutions is possible. A recent review covers standard separations of inorganic ions and metal complexes such as ion chromatography, as well as advanced techniques such as capillary electrophoresis (CE) [41]. The advantages of CE include the high separation speed (10 min) and separation efficiency, and the small sample volumes Met. Ions Life Sci. 2, 1–30 (2007)

6


required (several nL). Metals can be detected either by nonspecific UV detection, or by using an element-specific ICP-MS coupled to the CE instrument.

3.

ANCIENT AND MODERN USES OF NICKEL

The use of Ni can be traced back to the beginning of the Bronze Age in Mesopotamia in 3500 BC. The ancient bronzes produced in the area had a Ni content of up to 2%. The naturally occurring nickel–copper alloys were used by the Chinese during their Bronze Age from ca 2000 BC. However, the real understanding of Ni as an element dates to more contemporary times. In 1751 the Swedish chemist and metallurgist, Cronstedt isolated a white metal, which he called nickel, from a mineral originating from the Los Cu–Co mines in Sweden. It is not quite evident what was the Ni containing mineral, from which Ni was identified by Cronstedt, but very probably it was ‘kupfernickel’ now called niccolite (NiAs). The name ‘nickel’ originates from the old name of this mineral. Although kupfernickel looked like a Cu mineral, one could not extract any Cu from it. Therefore the Saxon miners had named the mineral according to a spiteful dwarf ‘Nickel’ who had apparently turned the Cu in the ore into a non-available form. Nickel is an important element in modern industry. The largest use by far is the manufacture of stainless steel, an alloy that consists of 8% nickel, 18% chromium and 74% iron. Other special steels can contain up to 60% of Ni. There are more than 3000 known Ni-containing alloys. Finely divided Ni is used as a catalyst for hydrogenating vegetable oils. Batteries contain Ni, especially the advanced Ni–Cd batteries of electric vehicles. Nickel can be electroplated onto other metals to form a protective coating on electronic components. Various Ni compounds are used as pigments [42].

4. 4.1.

SOURCES OF ATMOSPHERIC NICKEL Natural Sources

There are large amounts of natural Ni present in the atmosphere, derived from windblown dust, volcanic ashes, forest fires, meteoric dust and sea salt spray. On a global scale, windblown soil particles from eroded areas can account for 30–50% of natural Ni emissions. Volcanoes eject material from the Earth’s mantle and this source can also account for a significant portion, perhaps 40–50% of Ni, in the air [1]. Nickel, which has been incorporated into vegetation can be released and dispersed by forest fires. Therefore, the natural background concentration in the air varies according to local sources and specific climatic conditions. Estimates from natural emissions are still rather inaccurate. Pacyna and Pacyna [2] estimate that the natural emissions would be roughly one-third Met. Ions Life Sci. 2, 1–30 (2007)


7

(30 000 tons) of the anthropogenic emissions. However, locally the ratio of natural to anthropogenic emissions can be entirely different, e.g., in the neighborhood of volcanoes.

4.2.

Anthropogenic Sources

Important sources of Ni are present and past mining activities, including foundries and smelters and refineries; diffuse sources such as piping, constituents of industrial and commercial products, combustion by-products, waste disposal, and traffic [43]. However, the main anthropogenic source of Ni is the combustion of oil, emitting 86 110 tons of Ni in 1995, which is more than twice as high as the corresponding emissions in 1983 (40 833 tons) [2]. The increase can be explained by increased production of electricity and heat by oil combustion worldwide. Actually almost 90% of the global anthropogenic Ni emissions originate from oil combustion [2]. The worldwide emissions of Ni from non-ferrous metal production in 1995 was 8876 tons [2]. Historical records, derived from peat bog profiles or ice cores (see also Section 6), have shown that trace metal pollution started to increase at the beginning of the Second Industrial Revolution, between 1850 and 1900, when worldwide Ni emissions have been estimated to be close to 240 tons year1 [1]. Over a hundred years later, in 1983, the total Ni emissions were already 55 650 tons yr1 [2]. Even though the emissions of most metals have decreased since 1980s, global Ni emissions reached a level of 95 287 tons yr1 in 1995, largely due to the increase in oil combustion. However, at a local scale trends can be different, e.g., in Finland Ni emissions declined by 50% between the 1990 and 2000 [44]. The decline is supposed to result from the use of more efficient techniques and combustion processes together with better process control. Removal of Ni containing particles has also become more efficient owing to the installation of new sulfur removal systems [45]. The largest anthropogenic sources of atmospheric Ni emissions are estimated to be in Asia where emissions show an increasing trend, due to increasing industrialization and lower efficiency of emission control [2].

5.

DEPOSITION AND FATE OF ATMOSPHERIC NICKEL

5.1. Air Quality and Deposition in Polluted Areas versus Remote Areas Nickel is emitted into the atmosphere mainly in particulate matter dispersed by the wind and deposited by both dry and wet deposition. The deposition rate and the transport distance depend on the location, stack height, size of the particle, and meteorological conditions. Nickel from anthropogenic sources is mostly in Met. Ions Life Sci. 2, 1–30 (2007)

8


the form of oxides and sulfates of rather small particle size (mass median diameter about 1 µm) of which some 15–90% is soluble [43]. Windblown dusts may contain mineral species of Ni, which are often in the form of sulfides. The type of ore will govern the form of Ni emitted to the atmosphere from an ore-crushing plant [46]. The major part of the particles deposit in the vicinity of the source, but smaller particles are transported over longer distances. This is especially true in winter when the particles can remain suspended in the air for long periods of time. In the Arctic, air measurements show that concentration of Ni is higher in winter than in summer by more than one order of magnitude [47]. The Ni concentrations in ambient air show considerable variation. In a remote area in the Canadian Arctic levels of 0.38–0.62 ng m3 were recorded [48], whereas a value of 124 ng m3 was measured in the vicinity of a Ni smelter [49]. At a background site in northern Finland, a level of about 0.58 ng m3 as an annual average value, was recorded during 1996–1998 [50–52], while a background value in southwestern Finland was 1.14 ng m3 [53]. In a polluted area in southwestern Finland, in the vicinity of a Cu–Ni smelter, daily peak values as high as 44–230 ng m3 have been observed [54]. Ranges from 9 to 60 ng m3 have been reported for European cities [43], and from 110 to 180 ng m3 for heavily industrialized areas [55]. Moreover, in remote areas near volcanoes values as high as 330 ng m3, have been measured [56]. According to Nriagu [1] average concentrations of Ni in urban areas are 52 ng m3, at rural sites 4.3 ng m3, and at remote sites 0.8 ng m3. Deposition of heavy metals including Ni has been monitored systematically in Europe at background sites since the 1990s in the EMEP programme (http:// www.emep.int/index_facts.html). In 2001 the measured Ni deposition varied between 54.6 and 7732.7 µg m2 [57]. The highest deposition was recorded in Svanvik, Norway, at a site located near the Russian border, and under the influence of the Ni–Cu smelters in the Kola Peninsula. The lowest annual Ni deposition was recorded at Irafoss, Iceland. In another study in a polluted area near a Cu–Ni smelter in southwestern Finland, the average annual Ni deposition during 1993–1998 was 64 mg m2 at 0.5 km from the main stack [58].

5.2. Regional Indicator Surveys: The Use of Mosses, Lichens, Bark 5.2.1.

Mosses

The use of mosses as biomonitors of atmospheric pollution on a regional scale were started in the late 1960s in Sweden [59,60] and were gradually adapted by other Nordic countries [61–63]. Mosses receive most of their nutrients directly from the atmosphere, although the interaction with local litter and soil should not be underestimated [64]. The general uptake efficiency of different trace elements by mosses decreases in the order Pb Co,Cr Cu, Cd, Mo Ni, V Zn As Met. Ions Life Sci. 2, 1–30 (2007)


9

[65]. The glittering feather moss (Hylocomium splendens (Hedw.) B.S.G.) and the red-stemmed feather moss (Pleurozium schreberi (Brid.) Mitt.) are the most common species used in regional surveys in Nordic countries [66–68], even though they are known to be sensitive to pollution and do not grow, e.g., in severely Cu–Ni contaminated areas [69]. According to Rühling [70] Ni concentrations in mosses over large parts of northern Europe are generally less than 2 µg g1. In Finland, the Ni concentrations of a moss survey, which has been carried out in five-year intervals since 1985, have varied between 0.46 and 79.7 µg g1 [68]. The highest concentrations have occurred near point sources, such as a large Cu–Ni smelter in southwestern Finland. Higher maximal concentrations have been found in the Barents region, which covers the northwestern part of Russia (the Kola Peninsula), and the northern parts of Finland and Norway, including both heavily polluted industrial sites (Nikel, Zapoljarnii and Monchegorsk), and some of Europe’s most pristine wilderness areas. In the Barents area, Ni concentrations varied from 0.96 to 396 µg g1 and the median concentration was 5.39 µg g1 [42]. Studies conducted near Ni– Cu smelters of Sudbury (Ontario, Canada) and a Ni refinery in Port Colborne (Ontario, Canada) showed that up to 980–1500 µg Ni g1 could accumulate in transplanted mosses in 30-day exposure periods. Naturally growing mosses in Ontario have been reported to contain 1.3–81 µg g1 Ni [71].

5.2.2.

Lichens

Epiphytic lichens have been extensively used in monitoring as well. The use of lichens as bioindicators of air quality was discovered by the Finnish botanist Nylander in the 19th century, when he noticed that lichens occurred only in the most sheltered locations in the famous Jardins du Luxembourg in Paris [72]. The physiology of lichens makes them sensitive and also vulnerable to air pollutants, especially to high levels of sulfur dioxide (SO2). Therefore, epiphytic lichens are the first to disappear from the immediate surroundings of pollutant sources [72–74]. However, since they obtain their nutrients directly from aerial deposition [75,76], they are efficient in trapping atmospheric heavy metals in low or moderate pollution conditions [71,77,78]. The epiphytic lichen Hypogymnia physodes (L.) Nyl. is widely distributed and fairly resistant to impurities in the air [79], therefore it has been used commonly in regional surveys of heavy metal deposition [80–82]. In a regional survey in Finland, the average Ni concentration of H. physodes was 2.5 µg g1 [83]. In the Kola Peninsula (Russia) near Ni–Cu smelters, H. physodes lichens were transplanted for a 3-month period, at distances ranging from 6 to 150 km from the smelters [73]. The Ni concentration decreased in lichens with distance from the smelter, but at 6 km the Ni concentration was as high as 2780 µg g1 (at 12 km 610 µg Ni g1; at 18 km 200 µg Ni g1; at 40 km 110 µg Ni g1; at 150 km 12 µg Ni g1). Met. Ions Life Sci. 2, 1–30 (2007)

10


In a survey carried out with seven lichen species near the Sudbury smelters in Ontario (Canada) concentrations of Ni, Cu, Fe, and S correlated with each other [84]. Concentrations of Ni ranged between 10 and 37 µg g1, Stereocaulon paschale (L.) Hofm. and Umbicularia spp. in particular accumulating larger quantities of Ni. In a more recent study by Case [85], carried out in the vicinity of the Sudbury smelters, the Ni concentration of Cladina species was on average 24 µg g1 and at a background site ca 2 µg Ni g1. Backor and Fahselt [86] studied Cladonia pleurota (Flörke) Schaer. in a heavy-metal-rich site near a roasting bed as well as at a local background site in Sudbury. The average Ni concentration near the roasting bed was 212 µg g1 compared with the local background site 71.6 µg g1.

5.2.3.

Bark

The use of tree bark as an indicator of air pollution increased in popularity after it was found that the pH of bark generally correlates well with atmospheric SO2 concentrations [87]. Results from the Kola Peninsula, Russia, and northern Finland indicated extremely high Ni concentrations in the bark of Scots pine (Pinus sylvestris L.) growing close to the Ni–Cu smelters in Monchegorsk, with a maximum value of 303 µg Ni g1 [88]. The average Ni concentration of the Finnish study plots was only about 1 µg g1, and the range 0.05–6.64 µg Ni g1. In a Canadian study, where bark samples were collected from a variety of tree species growing under varying contaminant loads, considerable variation among species was observed. Coniferous species (white spruce, black spruce, tamarack, white cedar, and jack pine) usually contained the highest concentrations of Ni ranging from 2 to 61 µg Ni g1 [71], and the Ni concentrations of deciduous species varied from 1 to 37 µg g1. In a Nigerian study carried out beside a road the Ni concentrations in bark of two tree species (Azadirachta indica A. Juss. and Gmelina arborea Linn. Roxb.) were on average 13 µg g1 [89].

5.3. Nickel Concentration in Aquatic and Biotic Media Natural background levels of Ni in water are relatively low: in open ocean water 0.2–0.7 µg L1, and in fresh water systems generally less than 2 µg L1 [43]. In a Finnish study [90] median Ni concentrations in lakes were 0.38 µg L1 (n 153) and in stream waters 0.52 µg L1 (n 1165). It is typical that heavy metal concentrations are lower in the headwater lakes than in streamwaters, possibly because of particle scavenging. Nieminen et al. [91] compared Ni concentrations in the surface layers of a peat bog at a polluted site in the vicinity of Cu–Ni smelters to that of a peat bog in a remote area. The Ni concentrations were 100 times higher near the smelter (260–913 µg g1) compared with those of the remote site (3–14 µg g1). Mannio Met. Ions Life Sci. 2, 1–30 (2007)


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et al. [92] studied enrichment of Ni in lake waters in Finland. They found out that an enrichment factor (EF) value of 2 was exceeded in only three lakes in Northeast Finland, near the Russian border, and the Kola Peninsula Ni–Cu smelters. These findings suggested that the current Ni concentration was more than twice the historical Ni concentration in those same lakes. Corresponding Ni EF values of lakes in northern Scandinavia have been reported to be less than 2 [93,94]. Results of a study of various aqueous and biotic media in four remote sites in Finland indicate no bioaccumulation of Ni (Table 1), possibly a reflection of the generally low Ni concentration at all sites [95]. For example, the Ni concentrations of wood ant (Formica spp.) and the liver of shrew (Sorex spp.) were low compared with those found in moss, which should reflect the amount of atmospheric Ni deposition. Wood ant and shrew represent an important part of the ecosystem food chain. According to Tyler [96] the critical concentration value for Ni in mosses is 20 µg g1 and correspondingly in freshwater 25 µg L1 [97]. Hence, the Ni concentration of both mosses and surface water in the remote Finnish sites was clearly below these ecotoxicological lowest effect values (Table 1).

Table 1. The range of median Ni concentration in aquatic and biotic media in 1–4 remote catchments in Finland (1989–1996). a Ni Media

n

( µg L1)

Aquatic media Bulk precipitation Throughfall Stemflow Soil water Groundwater Lake water Streamwater

308 279 61 61 60 152 101

0.31– 0.5 1.20– 5.37 7.99–10.90 8.63–11.89 0.03– 0.09 0.11– 0.54 0.16– 0.46 µg g1 dry matter

Biotic media Feather moss Humus layer Peat 0–80 cm b Red wood ant Shrew (liver)

Range between catchments

8 72 16 53 85

1.9–3.5 5.7–8.4 2.1 0.4–0.5 0.1

a

Adapted and modified with permission from BER from Tables 3 and 4 in [95]. b Unpublished data from the Finnish Forest Research Institute.


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In a study conducted in an ultramafic (serpentine) soil, where a Ni hyperaccumulating plant (Alyssum pintodasilvae T. R. Dudley) was common, herbivores such as grasshoppers and predators such as spiders had significantly elevated concentrations of Ni [98]. In contrast, the concentrations of Cr and Co, which are also enriched in ultramafic soils, but are not accumulated by A. pintodasilvaea, were not elevated in the studied invertebrates. Therefore, the presence of Ni hyperaccumulating plants appears to enhance the transfer of Ni from soil to herbivore and carnivore trophic levels. However, with the exception of hyperaccumulator plants, Ni does not biomagnify in the terrestrial foodweb [99], thereby its toxicity to higher trophic levels appears to be unlikely.

5.4.

Nickel Fluxes in Forested Catchments

The atmospheric deposition of heavy metals is dominated by dry deposition of aerosols and particulates [100,101]. Forest canopies are particularly effective in trapping suspended heavy metal aerosols because of the high surface area for interception [102]. It is well known that the precipitation can wash many contaminants from the surfaces of vegetation. Less known is the fate of elements as the solution travels through the soil horizons and moves into streams. Some part of the elements absorbed by plants remain incorporated in stems, leaves, or moss tissues and are released back to the nutrient cycle only after senescence and litter decomposition. Input–output budgets at catchment and stand levels, therefore, are a useful means of describing the mobility, retention and fluxes of elements in the environment [103–108]. However, there are only a few elemental budgets published for Ni. Ukonmaanaho et al. [108] studied Ni input–output budgets both at plot (stand)- and catchmentscale in two remote boreal forest areas in southern (Valkea-Kotinen, Norway spruce dominated) and eastern (Hietajärvi, Scots pine dominated) Finland. Results of the plot-scale budgets showed that retention of Ni (by canopy, vegetation, soil) took place at both the Hietajärvi plots, whereas at the Valkea-Kotinen plot the Ni flux at 40 cm depth in soil was greater than the total inputs (TF LF) (Figure 2). Furthermore, if the internal fluxes (TF LF) are ignored, retention of the atmospheric Ni took place only at one of the two Hietajärvi plots. At the catchment scale, Ni outputs from Valkea-Kotinen and Hietajärvi were less than atmospheric inputs (Table 2), indicating that much of the atmospheric inputs were retained within the catchments. Most of the retention at Hietajärvi and Valkea-Kotinen was associated with the terrestrial part of the catchment, which includes retention in the tree canopy, as well as in soil and uptake and accumulation into plant biomass. As the stands at both Valkea-Kotinen and Hietajärvi are old growth forest, the net increment growth can be considered close to zero [109] and therefore net uptake of elements by trees is negligible. Assuming that the whole upland forested area of the catchments acts as indicated by the Met. Ions Life Sci. 2, 1–30 (2007)


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Figure 2. Plot-scale annual (1994–1996) average input fluxes and output budgets of Ni for: (a) Hietajärvi plot 1; (b) Hietajärvi plot 4; and (c) Valkea-Kotinen plot 1. Reprinted from [108] with permission from Elsevier.

plot scale budgets, i.e., Ni leaching atmospheric inputs of Ni (Figure 2b, c), most of the Ni retention in the terrestrial part of the catchment takes place below a depth of 40 cm in the soil profile. Retention of Ni2 can take place in B horizon, where cation-adsorbing Al and Fe hydroxides/oxides are enriched. There are Met. Ions Life Sci. 2, 1–30 (2007)

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Table 2. Mean annual (1994–1996) Ni input–output budgets, retention of atmospheric inputs and transfer of Ni in the Valkea-Kotinen and Hietajärvi catchments in remote areas in Finland.a

Catchment area (km2) Bulk deposition (mg m2 yr1) Total deposition (input) (mg m2 yr1) Runoff water (output) (mg m2 yr1) Output/input (%) Retention (input output) (mg m2 yr1) Relative retention (input output/input) (%) Deposition to terrestrial area (g yr1) Deposition to lake (g yr1) Terrestrial retention (%) Lake retention (%) Terrestrial transfer to lake (g yr1) a

Valkea-Kotinen

Hietajärvi

0.3 0.19 0.35 0.16 46 0.19 54 99 7 88 12 48

4.6 0.17 0.33 0.05 16 0.28 84 1370 144 85 15 285

Adapted with permission from Elsevier from [108].

considerable areas of peatlands in both catchments, which may be very important to the terrestrial retention observed. Sedimentation of organometal complexes is the main process by which heavy metals are retained within lakes [106,110,111].

6.

HISTORICAL RECORDS OF NICKEL DEPOSITION

6.1. Sediment The chronological geochemical record contained within sediments has been used worldwide to describe the history of contamination [112]. Lake sediments have not only shown the intensity of Ni pollution in the surroundings of the world’s largest Ni smelters at the Kola Peninsula in Russia [113] and at Sudbury in Canada [114] within the last century, but also the recent recovery of lakes reflected in the decreasing concentrations of Ni in the most recent layers. Lake sediments have also shown that the impact of these smelting activities is primarily localized rather than regional [115]. The release of Ni from industrial areas into the environment in the 1970s/1980s, and the recent decline is well recorded in delta systems, such as that of the Danube [116] and the Venice lagoon [117]. The slight enrichment of Ni in recent sediment layers of the Culiacan river estuary in Northwestern Mexico was related to population growth [118]. Studies in the UK have shown that even relatively remote sites have experienced enhanced atmospheric deposition of anthropogenically derived Ni for over Met. Ions Life Sci. 2, 1–30 (2007)


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100 years, and the contamination might have started before industrialisation [119]. Sediment records from a remote subalpine lake in Taiwan covering a time span of 2600 yr indicated that aeolian Asian dust particles have played a significant role in the elevated deposition of Ni during dry periods [120]. The natural enrichment of Ni (up to 0.1% Ni) in a lake sediment core from Finland was related to the prolonged erosion of metalliferous black shales during glacial retreat ca 9000 years ago [121].

6.2. Peat Bog Ombrotrophic peat cores have recently proved to be meaningful archives of recent as well as ancient atmospheric metal deposition, especially for Pb [122– 124] and Hg [125–128]. In contrast, there are only a few systematic studies on the distribution and fate of Ni in ombrotrophic bogs [129–134]. The results indicate that the fate of Ni is strongly dependent upon the mineralogical form in which it is originally deposited, but also on the geochemical and hydrological conditions in the peat layers, and the peat accumulation rate [133]. Due to the high deposition rate of Ni near a Cu–Ni smelter, recent smelter emissions are to some extent recorded in the peat profile [132]. In the long term, however, the bulk of deposited particles are expected to dissolve in the peat layers [134]. Even the emission of Ni sulfide particles from a nearby Ni mine was not well reflected in the peat profile, although these minerals had been expected to be chemically stable in the anoxic peat layers [132]. The crucial factor might be the early oxidation of particles in the aerated surface layers of the bog [133]. The retention of anthropogenic Ni from long-range atmospheric transport shows some (apparent) inconsistencies [129,130,132]. In a peat core taken at a low-background area in Finland, post-depositional processes could not be excluded [132]. In contrast, the deposition of dust-related Ni phases appeared to be well preserved in an age-dated Swiss peat core (Figure 3) [129]. The highest concentrations of Ni were found during the Younger Dryas cold climate event (centered at 10 590 yr B.P. according to 14C dating) when background values were exceeded by about 40 times. Elevated Ni accumulation rates were also found at 8230 and 5320 years B.P. according to 14C dating, which are consistent with the elevated dust fluxes recorded by Greenland ice cores. Silicate mineral particles are well preserved in acidic peat for thousands of years [135]. Apparently, when silicate phases are the main Ni-bearing host, Ni is well preserved in the peat profile.

6.3.

Polar Snow and Ice

Ice cores have provided much important information about temporal trends in atmospheric metal deposition in Greenland [137–139], the European Alps [140], and Antarctica [141]. However, to the best of our knowledge, there is only one Met. Ions Life Sci. 2, 1–30 (2007)

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Figure 3. Nickel and scandium concentrations in a Swiss peat core. The shaded area (ca. 5320–8230 years B.P. according to 14C dating) corresponds to the Holocene Climate Optimum when rates of atmospheric soil dust deposition were at their lowest [122,136]. Times of high dust deposition are indicated by elevated Sc concentrations in the peat [136]. Reprinted from [129] with permission from the American Chemical Society.

single study which includes Ni concentration data [142]. Nickel concentrations of a 12.5-m-long ice core from Devon Island, Canadian Arctic, ranged between 0.6 and 144 pg g1. The wide dispersion of concentration values is highly dependent on variations due to the strong seasonal influence on the mechanism of atmospheric transportation [142]. Met. Ions Life Sci. 2, 1–30 (2007)


7. 7.1.

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BIOAVAILABILITY AND MOBILITY OF NICKEL IN SOILS Uptake and Translocation of Nickel by Plants

Nickel is a natural constituent of plant tissues ranging normally from 0.05 to 10 µg g1 of dry matter [7,143]. The range in concentrations mainly reflects the differences between plant species in uptake and in root-to-shoot transport of Ni [143]. Although threshold concentrations for the lethal Ni toxicity are commonly reported to be less than 100 µg g1 [7,71,144–146], Ni concentrations as high as 600 µg g1, have been found in leaves of maples (Acer spp.) growing close to the Port Colborne Ni refinery in Ontario, Canada [147] and ⬃580 µg g1 in Scots pine needles of tree stands in the vicinity of a Cu–Ni smelter in Finland [148]. Such high values are mostly due to surface contamination of the field-grown plants by Ni-containing dust. However, it is also well known that certain endemic plant species growing on ultramafic soils, the so-called hyperaccumulators, can contain more than 1000 µg Ni g1 inside their tissues [149] (see also Chapter 2). In a Scots pine stand growing close to a Cu–Ni smelter, more than half of the Ni in precipitation originated from the wash-off of canopy-deposited dust [150] and pine needles exposed to aerial deposition contained tens of times more Ni than the needles protected from this source [148]. The measured Ni concentrations of boreal plant species growing naturally in Ni polluted forests are variable (Table 3), which may be, at least partly, due to varying degrees of surface contamination. Although root uptake is the main pathway for Ni access in higher plants, there is much evidence showing that Ni is also available to plants through the foliage both in conditions of excess Ni [153,154] and Ni deficiency [155]. Bryophytes and lichens are especially efficient in absorbing elements directly across their surfaces because they have neither epidermis or cuticle layer, nor roots [156]. Not surprisingly, therefore mosses and lichens are widely used as biomonitors of metals (see also Section 5.2). Most of the Ni deposited on foliar surfaces does not penetrate into the living cells through the cuticle of higher plants, and therefore does not have direct toxic effects on metabolism. Consequently, the Ni concentrations of field-grown plants are usually of little relevance in terms of a plant’s physiological response, even though they can be important in assessing risks to animals or humans. Washing of plant material prior to analysis is often recommended as a means to remove or separate the particulate materials contaminating plant surfaces [157]. However, in studies by Kozlov et al. [158], as well as by Nieminen et al. [148] only a minor part of surface-deposited Ni on tree foliage was removed by washing. The secretion of organic compounds by plant roots may affect the solubility of Ni in the rhizosphere soil [143,159]. There are several examples showing that the presence of other compounds in the substrate can either increase or decrease the Ni uptake, depending on their quantity, quality, and the characteristics of the plant in question. Nickel uptake by Scots pine seedlings grown in quartz-sand Met. Ions Life Sci. 2, 1–30 (2007)

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Table 3. Nickel concentration of some boreal plant species near the Cu–Ni smelter at Harjavalta (Finland), the Ni smelter at Monchegorsk (Kola Peninsula, Russia) and a background site in Finland. Ni ( µg g1)

Species Near the smelters Vaccinium myrtillus L.a Vaccinium vitis-idaea L.a Vaccinium vitis-idaea L.b Empetrum nigrum L.b Deschampsia flexuosa L. (Trin.) a Pinus sylvestris L., needlesc

1-year-old 2-year-old 3-year-old

959 945 22 54 1452 44 49 44

1-year-old 2-year-old 3-year-old

4.8 3.5 5.2 11.8 3.8 2.3 1.8

Background Vaccinium myrtillus L.d Vaccinium vitis-idaea L.d Empetrum nigrum L.d Deschampsia flexuosa L.(Trin.) d Pinus sylvestris L., needlese

a

2 km from the Ni smelter, Kola Peninsula, Russia [151]. 2 km from the Cu–Ni smelter, Harjavalta, Finland [152]. c 0.5 km from the Cu–Ni smelter, Harjavalta, Finland [150]. d Background value from Finland [152]. e Unpublished data from the Finnish Forest Research Institute. b

substrate was enhanced in the presence of copper [146], while Cataldo et al. [160] found that Ni absorption by hydroponically grown soybean plants (Glycine max (L.) Merr. cv. Williams) was completely inhibited in the presence of Cu. Singh et al. [161] observed that an increasing supply of urea appeared to mitigate the toxicity of Ni to wheat (Triticum aestivum L.). The presence of organic acids or inorganic ligands in soil solution results in formation of Ni complexes, which may either inhibit or enhance root uptake, depending on the characteristics of the Ni complex formed. For example, Molas [162] reports that Ni(II)-EDTA was less enriched and less toxic than Ni(II)-citrate or Ni(II)-Glu to hydroponically grown cabbage plants. Generally, the ionic form of Ni2 is taken up relatively easily by plants, but many chelated forms appear to be less available [160,163]. Highmolecular-weight solutes are prevented from entering the apoplasm of root cells by the diameter of pores [143]. Plants can restrict the apoplastic passage through the establishment of a suberin-rich transport barrier in the roots [164]. Met. Ions Life Sci. 2, 1–30 (2007)


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Although Ni is generally considered to be a readily mobile element in plants [8,143], the mobility within a plant is regulated by the formation of organic Ni complexes [165]. According to Tiffin [166] Ni is present in anionic organic complexes in the xylem of most plant species, which suggests that Ni is chelated by an organic ligand (carrier) before translocation to the shoot. Timperley et al. [167] reported that Ni in the foliage of several tree species was mainly complexed, and either in a neutral or positively charged species. In some plant species, e.g., oat (Avena sativa L.) and legume species (Fabaceae) associated with Rhizobium bacteria such as peas and beans, Ni is primarily enriched in the seeds [71,143]. Cataldo et al. [168] concluded that the leaves of soybean plants were a major sink during vegetative growth, while more than 70% of the Ni in shoot was remobilized to the seeds at senescence. During autumnal senescence, 5–20% of the Ni incorporated in the senescent needle mass was translocated to the remaining tissues of adult Scots pine trees growing in conditions of excess Ni [169].

7.2. Importance of Partitioning for Bioavailability and Mobility Total Ni contents in the soil might depict the potential availability of Ni, but in most cases the total content is of little relevance in terms of Ni availability. The basic aim of a number of soil extraction schemes is to estimate the metal pool available to plants or other soil biota [170]. Single extractants such as dilute acids (HCl, NH 4 , HNO3) or solutions containing chelating agents (DTPA or EDTA) are traditionally used to estimate the bioavailable proportion of soil metal [7]. However, when applied to soils with a wide range of soil characteristics, they fail to predict metal bioavailability adequately [171]. In addition to the mineralogical form of a metal, the key properties controlling its solubility and availability are soil solution pH, dissolved organic matter, and solid-phase metal oxide and organic matter content [172]. In general, the solubility of Ni is inversely related to the soil pH [8], albeit this dependence may be affected by organic complexation. Poulsen and Hansen [173] examined the influence of citrate and arginine on Ni sorption to a sandy loam soil and its dependence on pH. As the sorption edges for Ni without ligands showed 50% sorption at approximately pH 5, the presence of citrate depressed sorption, with the 50% sorption shifting to a pH value of more than 7.5. In contrast, arginine had almost no influence on sorption over the entire pH range studied. Poulsen and Hansen [173] concluded that the trivalent nickel–arginine complexes (NiH3arg23) were sorbed to cation exchange sites, while sites for the bonding of monovalent negatively charged nickel–citrate complexes (Nicit) were sparse. According to Wolt [174] and Kabata-Pendias [8], humic and fulvic acids, as well as simple carboxylic acids derived from the decomposition of organic matter, are sufficiently abundant in soil solutions and form complexes with Ni of sufficient Met. Ions Life Sci. 2, 1–30 (2007)

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stability, that these can have comparable effects on Ni sorption as citrate. These processes might explain the mobility of Ni in podzolic soils impacted by atmospheric Ni pollution [175]. The aim of the so-called sequential extraction techniques [176,177] is to dissolve various solid fractions step by step and to liberate the metals associated with each fraction, starting with the most accessible fractions and sequentially dissolving the increasingly recalcitrant fractions by using increasingly aggressive reagents. According to the results of sequential extractions most of the Ni in soils should be associated with iron and manganese oxides (reducible fraction), as well as with the residual fraction [178–180]. However, the reagents used in these sequential techniques can never affect only a single fraction, and these results have always to be interpreted with caution. Furthermore, there is no evidence indicating which solid-phase fraction correlates best with uptake by organisms [171]. Element partitioning may be further modified in the rhizosphere and within the digestive tract of soil-dwelling organisms [172]. In the case of soil invertebrates, extractant methods that attempt to mimic their digestive processes would be the most suitable ones to predict the availability of soil Ni to them. Since plants access metals in the soil principally through the soil solution, it might be reasonably expected that the determination of Ni concentrations in the soil solution would provide the best predictor of Ni availability to plants. However, the prediction capacity of soil solution concentrations is not yet clear. Although the relatively high Ni concentration in soil percolation water was interpreted as reflecting higher downward leaching of Ni compared with Cu in forest soils adjacent to Cu–Ni smelters [175], Ni uptake by Scots pine roots did not appear to be more efficient than that of Cu [146,181]. In most Ni exposure experiments in which plants have been grown hydroponically, the Ni concentrations are often so high that the results do not have any ecological relevance. Toxic threshold concentrations of nutrient solutions reported by Mishra and Kar [163] vary from 0.5 to 300 mg L1 depending on the plant species in question, whereas Ni concentrations in soil solutions taken from a dead Scots pine stand next the Ni–Cu smelters at Monchegorsk in the Kola Peninsula (Russia) ranged from 0.1 to 2.9 mg L1 [3]. The total Ni concentration (dry ashing and HCl extraction) in the topmost organic layer of the same pine stand was ca. 3000 µg g1 of organic matter [182] and ⬃70 µg g1 of dry matter in the mineral soil horizons [183]. An average value of 20 mg kg1 for Ni of world soils, and a range of 5–500 mg Ni kg1, have been reported by Adriano [7]. However, in ultramafic soils, which are naturally enriched by Ni, values ranging from 100 to 3000 mg Ni kg1 have been reported in the UK and Scandinavia [184]. According to Tamminen [185] the median concentration of Ni in the organic layer of upland forest soil in Finland is 8.2 µg g1 of dry matter.



8.

21

SUMMARY AND CONCLUSIONS

During the last century Ni concentrations in soils and sediments increased worldwide. Nickel is frequently enriched in oil and coal, and oil combustion is currently the largest source of atmospheric Ni by far, although Ni emissions from mining and smelting activities have caused extensive damage in the areas surrounding large industrial complexes. Nickel is a valuable metal for modern industry, mostly due to its expanding need as a crucial component of stainless steel. The environmental mobility of Ni is considered to be low under neutral to alkaline and reducing conditions, but in acidic organic rich soils where Ni can be quite mobile, Ni contamination may pose a risk to groundwater quality. Even though the ionic form of Ni2 is taken up relatively easily by plants, chelated highmolecular-weight compounds are less available. Therefore, Ni does not appear to be subjected to biomagnification in terrestrial food webs, with the exception of the Ni hyperaccumulating plants. However, frequent application of sewage sludge on agricultural fields may result in substantial Ni absorption by some crop plants. Although Ni is an essential minor element, it is toxic in excess doses. Until now there has been more concern about the toxicity than possible deficiencies of Ni. Nevertheless, knowledge of the mechanisms underlying both the deficiency and toxicity of Ni have remained relatively limited. The content of the following Chapters of this book is intended to help fill these gaps and to increase our understanding of the relationships between Ni and organisms.

ABBREVIATIONS AAS AFS AMAP BC B/C-horizon BD BP CE DTPA EDTA EF Eh EMEP EMPA EPA

atomic absorbtion spectrometry atomic fluorescence spectrometry Arctic Monitoring and Assessment Program before Christ mineral and parent material horizon bulk deposition before present capillary electrophoresis diethylenetriaminepentaacetate ethylenediamine-N,N,Nⴕ,Nⴕ-tetraacetatic acid enrichment factor standard potential Co-operative Programme for Monitoring and Evaluation of the Long-Range Transmission of Pollutants in Europe electron microprobe analyzers Environmental Protection Agency


22

ICP-MS ICP-OES LF LOD TF TD UV WHO WQC XAFS XANES XRF


inductively coupled plasma mass spectrometry inductively coupled plasma optical emission spectrometry litterfall limit of detection throughfall total deposition ultraviolet radiation World Health Organization water quality criteria X-ray fine-structure analysis X-ray absorbtion near-edge structure X-ray fluorescence spectrometry

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2 Nickel in the Environment and Its Role in the Metabolism of Plants and Cyanobacteria Hendrik Küpper and Peter M. H. Kroneck Fachbereich Biologie, Universität Konstanz, D-78457 Konstanz, Germany

1.

INTRODUCTION 1.1. Coordination Chemistry of Nickel 1.2. Nickel: The Catalyst of Early Life 2. NICKEL AS A MICRONUTRIENT FOR PLANTS AND CYANOBACTERIA 2.1. Nickel Biochemistry and Its Physiological Role in Plant Metabolism 2.2. Nickel Deficiency 3. NICKEL AS AN ENVIRONMENTAL POLLUTANT AND ITS EFFECTS ON PLANTS 3.1. Nickel Toxicity 3.2. Sources and Occurrence of Nickel Pollution 3.3. Mechanisms of Nickel-Induced Inhibition of Plant Growth 3.3.1. Inhibition of Root Function 3.3.2. Inhibition of Photosynthesis 3.3.3. Oxidative Stress 3.3.4. Other Effects 3.4. Mechanisms of Resistance Against Nickel Toxicity 3.4.1. Exclusion and Sequestration 3.4.2. Binding by Strong Ligands 3.4.3. Other Mechanisms Metal Ions in Life Sciences, Volume 2 © 2007 John Wiley & Sons, Ltd


32 33 35 37 37 39 40 40 41 42 42 43 44 45 45 45 46 47

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48 49 50

NICKEL HYPERACCUMULATION 4.1. Biological Function of Nickel Hyperaccumulation 4.2. Mechanisms of Nickel Hyperaccumulation 4.3. Use of Nickel Hyperaccumulator Plants for Phytomining and Phytoremediation 5. OUTLOOK ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

1.

52 53 54 54 55

INTRODUCTION

Many heavy metals such as copper, nickel, and zinc are well known as essential trace elements for plants and cyanobacteria [1,2] (Figure 1). Productivity of phytoplankton in the oceans is often limited by the availability of heavy metals, not just Fe as some people tend to think, but also for example Ni [3]. In the first part of this chapter we will focus on the biochemistry of nickel, followed by its beneficial aspects and essential functions of metals in the metabolism of

Figure 1.

Scheme of the dose–response relationship for metals in plants.


Ni METABOLISM IN PLANTS AND CYANOBACTERIA

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plants and cyanobacteria. Furthermore, we will summarize recent developments concerning the uptake of Ni into plants and cyanobacteria and the effects of its deficiency on their metabolism. However, nickel can also be toxic to plants and cyanobacteria. Above the threshold leading to growth inhibition (Figure 1), a variety of toxic effects have been observed in all species, the relative importance of which depends on the concentration of the metal, the plant species, and different environmental conditions, including irradiance and soil conditions. General aspects of such inhibitions have been discussed by us recently [4], here we will concentrate on nickel-specific effects, including relations between nickel and other heavy metals. Plants have developed a number of strategies to resist the toxicity of heavy metals, as reviewed, e.g., by Prasad and Hagemeyer [5] and Küpper and Kroneck [4]. However, many of these mechanisms have been investigated with other metals, and it is unclear how far they also apply to nickel resistance. Plants that actively prevent metal accumulation inside the cells are called excluders, these represent the majority of metal-resistant plants [6]. Other resistant plants deal with potentially toxic metals in just the opposite way, i.e., they actively take up metals and accumulate them. These plants, which have been named ‘hyperaccumulators’ [7], are able to accumulate several percent of metals in the dry weight of their aboveground parts. The active accumulation in the aboveground parts of hyperaccumulator plants provides a promising approach for both cleaning anthropogenically contaminated soils (phytoremediation) and for commercial extraction (phytomining) of metals from naturally metal-rich (serpentine) soils [8,9]. Previously we reviewed general aspects of hyperaccumulation [4], in this contribution we will summarize some most recent findings concerning the metal hyperaccumulation trait in plants, and nickel-specific aspects as nickel is the metal for which phytomining is closest to commercial application. Research on the biological chemistry of nickel has become an important issue, as documented by a remarkable number of publications in biology, ecology and chemistry. Several nickel-dependent enzymes reveal complex multi-metal catalytic sites with unusual chemical and spectroscopic properties, which have attracted the interest of chemists and physicists. In view of the data accumulating in the field, we recommend two books for introduction: (i) The Bioinorganic Chemistry of Nickel [10], and (ii) Biochemistry of Nickel [11]. Furthermore, the Handbook on Metalloproteins [12] and the Handbook of Metalloproteins [13] summarize valuable information on structural and biochemical properties of nickel enzymes. The comprehensive reviews [2,14–24] are also recommended for further reading.

1.1.

Coordination Chemistry of Nickel

If there ever was a utilitarian metal, it is most likely nickel, which was discovered in 1751 by the Swedish chemist A. F. Cronstedt. The name comes from the Met. Ions Life Sci. 2, 31–62 (2007)

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German ‘Kupfernickel’, loosely meaning Devil’s copper. This transition element has a broad range of applications in daily life, it is used in everything from coins to automobiles to jewellery, and new uses are discovered all the time. Nickel is an excellent catalyst for many reactions. It is used for a large number of industrial and research applications, by itself, or in combination with other metals. Perhaps the most famous catalyst is Raney nickel, developed in 1920. There are parallels between the biological and industrial chemistries of nickel. Three of the prominent nickel enzymes, i.e., hydrogenase, carbon monoxide dehydrogenase, and methyl-coenzyme M reductase, catalyze reactions which correspond to industrial processes, namely hydrogenation, desulfurization, and carbonylation [18,21,25]. Positioned between cobalt and copper in the periodic table of the elements, nickel has the electronic configuration [Ar]3d84s2. The stable isotope 61Ni (1.14% natural abundance) has been employed successfully to study nickel-dependent enzymes by electron paramagnetic resonance (EPR) spectroscopy because of its nuclear spin I ⫽ 3/2 which gives rise to a characteristic hyperfine splitting into four lines [26]. The coordination chemistry of nickel encompasses a wide variety of geometries, coordination numbers, and oxidation states [27–29]. In aqueous solution, the common oxidation state is Ni(II), of configuration 3d8. This state is associated mainly with octahedral or tetragonally distorted octahedral geometry, but square planar (diamagnetic, S ⫽ 0) and tetrahedral (high-spin, S ⫽ 1) geometries are well known. Square pyramidal and trigonal bipyramidal Ni(II) complexes have also been reported. Examples of the more unusual oxidation states, Ni(⫺I, 0, ⫹I, ⫹III, ⫹IV) are comparatively sparse. However, in biology, both Ni(I) (3d9) and Ni(III) (3d7) have been observed and characterized by spectroscopic methods and X-ray crystallography. Usually, the Ni(I) state is associated with distorted octahedral or square planar geometry, whereas Ni(III) is usually low-spin, with distorted octahedral and, less frequently, trigonal bipyramidal geometry. Changes in the oxidation state of nickel during the catalytic cycle of an enzyme may involve substantial changes in the geometry around the metal center, a process that will be crucially dependent on the flexibility of the biological ligand. Changes in the spin state and size of the nickel ion have also to be accommodated. Note that in the case of methyl-CoM reductase, nickel is coordinated by a unique macrocyclic ligand, the so-called corphin, which appears to accommodate these changes well, and also that even in the highest oxidation state found for nickel in biology, Ni(III), there is still a high number of d electrons available [24]. Nickel has special parallels with cobalt in biological chemistry, but it also appears reasonable to compare nickel with its other immediate neighbors in the sequence Fe, Co, Ni, and Cu. Moving from left to right, there will be a change towards greater preference for nitrogen and sulfur ligands over oxygen donors as the chemical softness of the metal increases. In addition, the size of the divalent cation decreases, at least partly explaining the change in macrocycle associated Met. Ions Life Sci. 2, 31–62 (2007)


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with Fe, Co, and Ni. The porphyrin ring is highly conjugated and less flexible, while the F430 macrocycle is a saturated, flexible nonplanar ring which effectively complexes the smaller Ni(II), but allows for spin-state and oxidation-state changes affecting the Ni ionic radius [30–33]. Oxidation state (⫹II) is the common stable oxidation state for all four metals, but the relative stability of the (⫹III) oxidation state decreases and that of the (⫹I) oxidation state increases on moving from Fe to Cu. The occupancy of the 3d orbitals increases and this is emphasized by the increasing stability of the (⫹I) oxidation state. These electrons, particularly in the lower oxidation states, will be exposed as they occupy σ orbitals [25,34]. Ligand exchange reactions at Ni(II) centers are relatively slow, explaining why Ni(II) is unable to catalyze enzymatic reactions in place of Mg(II), and suggesting that Ni(II) may not be particularly favored for biocatalysis [15]. On the other hand, there is the unique example of urease, where nickel functions as a Lewis acid, a role that is usually assigned to zinc. Thus, the use of nickel by organisms, despite the requirements for providing specific pathways for transport and mechanisms for insertion into sites, strongly suggests that the metal confers substantial additional catalytic efficiency [16,35,36]. For example, Helicobacter pylori which is a well-established etiologic agent of gastritis, produces up to 6% of its soluble cell protein in form of the nickel-enzyme urease [37]. The ammonia produced by this multimeric enzyme has been postulated to allow the bacterium to survive and colonize the low pH environment of the gastric mucosa. The NixA nickel transport protein has an extremely high affinity (KT ⫽ 11.3 nM) import mechanism well suited for scavenging Ni2⫹ from the low concentrations found in human serum and presumably in the gastric mucosa [38].

1.2. Nickel: The Catalyst of Early Life The environment in which life began was most likely electron-rich, and gases such as dihydrogen and carbon dioxide, and metal sulfides were available [39–41]. Thus, special catalysts were needed to handle these gases, and early life-forms featured a rich biochemical role for nickel [42]. In some sense it looks as if nickel, similar to cobalt, was most useful when metabolism was based on such chemicals, but after the advent of dioxygen their value diminished [43]. One possible candidate for an early nickel catalyst could be the mineral greigite [Fe5NiS8], an iron–nickel sulfide. Its molecular structure is very similar to that of the thiocubane [4Fe–4S] unit of the ferredoxins, as well as to the cuboidal complex in the active site of the enzyme acetyl-coenzyme A synthase/carbon monoxide dehydrogenase [19,44]. Another important piece of information along these lines comes from the observation that a freshly precipitated aqueous slurry of coprecipitated nickel and iron sulfides could produce acetate in the presence of carbon monoxide and methyl sulfide, CH3SH [45]. Note that the NiFe hydrogenases which catalyze the heterolytic cleavage of molecular hydrogen, according to the Met. Ions Life Sci. 2, 31–62 (2007)

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Figure 2. Schematic representation of the nickel centers (grey) in (A) NiFe hydrogenase from Desulfovibrio gigas (PDB file 2FRV; Ni coordinated to Cys-68, 65, 530, 533), and (B) carbon monoxide dehydrogenase from Carboxythermus hydrogenoformans (PDB file 1SU7; Ni coordinated to Cys-256 and three bridging sulfide anions). Met. Ions Life Sci. 2, 31–62 (2007)


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reaction H2 L H⫺ ⫹ H⫹ L 2H⫹ ⫹ 2e⫺, are the only enzymes known so far that carry both CO and CN⫺ in their active site [19]. To date, the most intensively studied enzymes that require nickel for catalysis number six, including urease, superoxide dismutase, the NiFe hydrogenases, methyl coenzyme M reductase, carbon monoxide dehydrogenase, and acetylcoenzyme A synthase [46]. Less well characterized are the acireductone dioxygenase [47–49] (see Chapter 12 of this volume) and glyoxalase I [50,51] (see Chapter 11 of this volume). In addition, there are specific proteins required for delivery and assembly of the nickel ion into the active sites of nickel-dependent enzymes [16,52,53]. In the case of Bradyrhizobium japonicum, for example, a single protein, HypB or nickelin, has recently been purified. It was shown to have two roles: (i) that of nickel binding and storage, with this function being dependent on the histidine-rich N-terminus; and (ii) that of hydrogenase expression which may require the nucleotide-binding motif and GTP hydrolysis [54]. In higher organisms the activity of nickel is confined mainly to one curious enzyme, urease, although the symbiotic anaerobic bacteria of higher organisms still use nickel in some dihydrogen reactions. Methanogens have kept their nickel hydrogenases and other nickel enzymes, but the methanogens belong to a special class of bacteria, the Archaea, most of which have been relegated to anaerobic niches within the geochemical world. They are thought to be of primitive ancestry and also use special cofactors, including the F430 ring chelate of nickel as well as a special sulfhydryl coenzyme, the so-called coenzyme M [55].

2. NICKEL AS A MICRONUTRIENT FOR PLANTS AND CYANOBACTERIA 2.1. Nickel Biochemistry and Its Physiological Role in Plant Metabolism Nickel was one of the last metals found to be essential for the growth of plants. In 1975 it was found to be the active center of urease [56,57], an enzyme that converts urea to ammonia and carbon dioxide, both of which can be re-used in plant metabolism. Urease itself and its function has been known for a very long time; it was the very first enzyme to be purified and crystallized in 1926 by James Sumner at Cornell [58]. He isolated and crystallized urease from Jack bean (Canavalia ensiformis) over a period of nine years. This work was rewarded with the Nobel prize in Chemistry in 1946. However, it took half a century before Ni was found to be its active center, end even now only a few studies have shown that Ni is an essential plant micronutrient sensu strictu [59–61]. The first crystal structure of urease became available in 1995, using recombinant enzyme from Klebsiella aerogenes [62]. In their original catalytic mechanism for urease Zerner and colleagues [63] proposed a two-nickel site, with the metals within a Met. Ions Life Sci. 2, 31–62 (2007)

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distance of 6 Å. Hereby, the oxygen of the urea carbonyl coordinates to one nickel with additional activation provided by a carboxylate. This mechanism could be extensively refined on the basis of several crystal structures while retaining many of its basic features (reviewed by Hausinger and Karplus, [64]), and will be dealt with in more detail in Chapter 6. Another function of Ni apart from urease activity is found in N2-fixing legumes. The symbiotic N2-fixing bacteria of these plants require Ni as the active center of hydrogenase, an enzyme that recycles H2 produced by a side reaction of nitrogenase [65]. For this reason, legumes (reviewed by Gerendás et al. [66]) and some diazotrophic cyanobacteria [67] have been reported to have an elevated nickel requirement. The structural and mechanistic details of nickel-containing hydrogenases will be dealt with in more detail in Chapter 7. Little is known about the transport of nickel into the plant and translocation within the plant. The only Ni(II)-specific transporter found so far (TgMTP1t2) seems to be involved in the hyperaccumulation of Ni in Thlaspi goesingense [68]. It was shown to be an alternative splicing product of an mRNA that can also produce a transporter (TgMTP1t1) for the divalent ions of Cd, Zn, and Co, but the protein itself has not been isolated and characterized. Some activity for Ni uptake was also shown for the Mg transporter AtMGT1 [69]. Genes for the family of nickel/cobalt transporters (NiCoTs) have been found in plants [70]. According to their sequence, these proteins have eight transmembrane helices with a highly conserved region in helix 2. They have been shown to mediate high-affinity uptake of cobalt and nickel in various bacteria and yeast [70–72]. But neither the function of the gene products (proteins) in plants has been analyzed, nor has any member of this family been isolated as a protein [70]. It has been postulated that nicotianamine, a nonproteogenic amino acid known to be involved in the transport of iron and copper in plants, is involved in nickel transport as well [73]. But this was only based on in vitro studies of binding constants, no physiological measurement in this direction has been carried out so far. In cyanobacteria, nickel appears to have a lot more functions than in eukaryotic plants. This is in accordance with the idea that cyanobacteria have evolved in a sulfidic environment [41]. So far, most studies on nickel metabolism in cyanobacteria have been directed towards the role of nickel in hydrogenase and hydrogen cycling [2]. Two distinct NiFe hydrogenases have been described in cyanobacteria: an uptake hydrogenase catalyzing the oxidation of dihydrogen H2 → 2H⫹ ⫹ 2e⫺) and a bidirectional hydrogenase which can either take up or produce dihydrogen [74–77]. The physiological role of the uptake hydrogenase appears to be coupled to nitrogen fixation [78]. Daday et al. [79] studied the effect of nickel on hydrogen metabolism and nitrogen fixation in Anabaena cylindrica. Throughout, nickel-containing cells had an active hydrogen uptake capacity, whereas nickeldepleted cells (⬍ 1.7 nM) had not. Note that no advantage of increased hydrogenase activity to the organism in terms of growth and nitrogen fixation could be observed. Under the conditions tested, neither hydrogenase nor urease were Met. Ions Life Sci. 2, 31–62 (2007)


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in any way essential for cell growth of A. cylindrica. Possibly, the significance of these enzymes for the organism becomes apparent only under certain stress conditions [80]. As hydrogen gas is seen as one future energy carrier by virtue of the fact that it does not evolve the greenhouse gas CO2, biological hydrogen production appears to have several advantages over other conventional hydrogen production processes. The hydrogen production by cyanobacteria offers some promise in this respect, although more research is needed before this commodity can be effectively utilized [81]. An interesting development came recently from the identification of open reading frames in the genomes of marine cyanobacteria with significant similarity to Nidependent superoxide dismutases (NiSOD), and the in vivo production of an active nickel superoxide dismutase from Prochlorococcus marinus MIT9313 in Escherichia coli [82]. Predicted secondary structures are consistent with four-helix bundles, and the N-terminal processing and metal coordination sites are fully conserved. The first NiSOD was found in Streptomyces species in 1996 [83]. NiSOD is a homohexameric enzyme. The monomers have a four-helix bundle structure and contain one Ni bound to an N-terminal nickel hook. The Ni ion is coordinated by His-1 and Cys-2 and Cys6. The geometry and the oxidation state cycle between square planar Ni(II), with the amino group nitrogen of His-1, the backbone nitrogen of Cys-2, and the thiolate sulfurs of Cys-2 and Cys-6 as metal ligands, and square pyramidal Ni(III) where the imidazole side chain of His-1 is the fifth ligand [84,85]. Mechanisms of nickel uptake in cyanobacteria are, however, even less known than those in higher plants although an early study of Campbell and Smith [86] of Ni2⫹ uptake in Anabaena cylindrica had shown very interesting results. By using radioactive 63Ni, these authors could clearly show that Ni2⫹ uptake in A. cylindrica was mediated by a highly Ni-specific, high-affinity (apparent KM ⫽ 17 nM) energy-dependent (i.e., active) uptake system. However, the protein mediating this uptake has still not been isolated. A gene cluster involved in nickel sensing and possibly nickel uptake was identified in Synechocystis [87], but apart from its sequence so far nothing is known about the physiological function and its biochemical mechanism for any of these genes. The sequence of the gene nrsD suggests that it may be a Ni2⫹ permease with a carboxy-terminal metal binding domain. As in higher plants, also in cyanobacteria NiCoT genes for putative nickel transporters exist, but no data are available concerning their function [70].

2.2.

Nickel Deficiency

Like most metals, nickel is in short supply in the oceans; Ni deficiency has been shown to limit the use of urea by phytoplankton [88]. Probably for this reason, some green algae have an alternative urea-degrading system, the ATP-dependent urea amidohydrolase–allophanate pathway [89]. Met. Ions Life Sci. 2, 31–62 (2007)

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In most plant species, nickel deficiency is rarely observed because only very minute amounts of this metal are needed for normal metabolism, and the adequate range between limiting and toxic concentrations is exceptionally large compared to other heavy metals (Figure 1; reviewed in detail by Welch [1] and Gerendás et al. [66]). For this reason, it took a long time before Ni was identified as being essential for plant growth (see above), and nickel has been termed an ‘ultramicronutrient’ [90]. However, a recent study [91] on Scots pine (Pinus sylvestris) showed an increased growth after addition of 20 mg nickel per liter of soil, indicating that nickel deficiency may be more common than previously thought. Symptoms of Ni deficiency are a consequence of the lack of urease, which leads both to an accumulation of urea to toxic levels and to a lack of usable nitrogen sources. Visible symptoms of urea toxicity are leaf tip necroses [59,60], while symptoms of nitrogen deficiency are interveinal necrosis and patchy necrosis of younger leaves [92]. In addition, it was reported that the disturbance of nitrogen metabolism, even when nitrogen is supplied as nitrate, leads to strongly reduced levels of alanine and thereby impaired protein synthesis [93]. Interestingly, urease activity seems not to be regulated apart from the limitation by Ni availability; as long as Ni is available, urease is expressed constitutively [94]. In contrast to all ‘normal’ plants referred to so far, plants that hyperaccumulate nickel can easily suffer from nickel deficiency. In a study of three Ni hyperaccumulating Brassicaceae, Küpper et al. [95] observed that these plants show severe symptoms of nickel deficiency on normal soil, and only an addition of several hundred ppm Ni(II) leads to normal growth. In addition to its function in urease, in hyperaccumulators nickel is used for deterring pathogens and herbivores for which it is predominantly stored in epidermal vacuoles (see below). Most likely the mechanism for Ni sequestration into these vacuoles is so efficient that it easily leaves all other parts of the plants Ni-deficient.

3.

NICKEL AS AN ENVIRONMENTAL POLLUTANT AND ITS EFFECTS ON PLANTS

Elevated concentrations of nickel inhibit plant metabolism, leading to various effects, depending on the type of affected plant and the environmental conditions during the stress. Recent general reviews on this theme include the book of Prasad and Hagemeyer [5] concerning higher plants, our review in MIBS-46 [4] concerning higher plants, algae, and cyanobacteria, and a review by Bertrand and Poirier [96].

3.1. Nickel Toxicity The threshold concentration of nickel that leads to toxicity strongly depends on the type of plant under investigation, because plants differ drastically in their Met. Ions Life Sci. 2, 31–62 (2007)


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ability to deal with nickel toxicity. Furthermore, toxicity thresholds for nickel depend, as for any other toxic metal, on the growth conditions including irradiance, pH, and speciation of the metal. For the charophyte Nitellopsis, for example, a 4-day LC50 of 290 µM was found [97]. For the submerged water macrophytic plant Elodea, a 7-day lethal concentration of 200 µM was reported [98], and for the floating macrophytic plant Spirodela a 14-day EC50 of around 80 µM was observed [99]. Cyanobacterial biofilms seem to be more sensitive; inhibition was observed already at about 9 µM Ni2⫹ [100]. Subtle changes in physiology, which may still be significant for survival in the natural environment, may occur at much lower concentrations. For example, a break-up of colonies of the floating higher aquatic plant Lemna was observed already at 5 µM Ni2⫹ (or 0.2 µM Cu2⫹) within 6 h of exposure [101]. And a 15% decrease of root growth was observed at 2.5 µM Ni2⫹ in a particularly sensitive strain of Silene paradoxa growing on nutrient solution in sand culture [102]. For terrestrial higher plants, toxicity thresholds are very hard to predict because often only a small fraction of the total metal in the soil is bioavailable, and the bioavailability varies by orders of magnitude, depending on the ingredients and pH of the soil. Complexation by weak ligands, such as carboxyl groups of organic acids, hardly affects metal uptake into the plants, but complexation by stronger ligands (simulated, e.g., by EDTA addition) decreases nickel uptake and thus toxicity [103]. When considering the impacts of nickel pollution on plants one should keep in mind that nickel is much less toxic to plants than other important environmental contaminants, in particular copper. For submerged waterplants, for example, nickel is more than hundred times less toxic than copper [97,98]. This is particularly important since nickel pollution in most cases occurs together with copper pollution, as described in more detail below. Furthermore, some plant species seem to be able to adapt to heavy metal toxicity very efficiently. An investigation on the effect of nickel pollution on the reproduction of birch did not find adverse effects in an area that had been heavily polluted by a smelter [104]. Similarly, a Ni-tolerant strain of Scenedesmus acutus could be derived from a sensitive strain by selection in Ni-rich media in the laboratory [105].

3.2.

Sources and Occurrence of Nickel Pollution

Anthropogenic nickel pollution originates from three major sources: smelters, mines, and municipal waste. Concentrations of nickel in freshwater ecosystems affected by mining operations vary by several orders of magnitude, depending on the way the wastewater is treated. Concentrations reported for a lake in Finland [106] usually range from 0.5 to 1.4 µM with a maximum around 4 µM. Similar data were obtained for polluted river water in China (max. 3.4 µM Ni [107]) and Canada (max. 7.3 µM Ni [108]). Mandal et al. [108] reported inhibition of algae under such conditions, Met. Ions Life Sci. 2, 31–62 (2007)

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but again the toxicity originated most likely from copper, which reached close to 1 µM concentrations that can be lethal to aquatic plants [98]. It is important to note, however, that most (up to 95%) of the Ni2⫹ in the water samples studied by Mandal et al. [108] was not bound to strong ligands, making it highly bioavailable. The worst heavy-metal-emitting smelter in Europe is the Severonikel nickel– copper smelter at Monchegorsk (Kola Peninsula, Russia), releasing 1.21 ⫻ 106 kg Ni and 0.83 ⫻ 106 kg Cu into the environment in 2001 [109] and causing severe damage to the environment in an area of more than 10 000 km2. Most of this damage, however, is most likely not due to Ni, but rather Cu, combined with soil acidification by acidic rain. A similar type of contamination occurs around other smelters, such as the Ni smelter at Coniston (Ontario, Canada), where the soil contains up to 1500 ppm Ni and 1300 ppm Cu at pH levels around 3.7 [110], or the Ni refinery at Port Colborne (Ontario, Canada), where up to 2900 ppm Ni are found in the soil near the smelter and over 200 ppm in an area of 29 km2 around [111]. Municipal waste can be another important source of nickel pollution. According to a recent study [112], Ni concentrations in municipal waste leachate can be in the range of tens of µM. The adverse effect of applying such leachates on rice fields are, however, most likely not due to nickel but to copper, which accumulates in the soil to similar concentrations as nickel but is much more toxic (see above).

3.3. Mechanisms of Nickel-Induced Inhibition of Plant Growth Among the mechanisms proposed to contribute to heavy metal damage are inhibition of enzymes, photosynthesis, oxidative stress, changes in lipid metabolism and disturbances in the uptake of essential microelements, as reviewed in general by Küpper and Kroneck [4]. However, many of the effects were examined only in vitro, some of them either could not be observed to operate in vivo [113] or could not be confirmed by in vitro studies, and in many cases alternative explanations for the effects observed may be more likely. In many studies, heavy metals were applied at extremely high concentrations, which are far from being ecologically relevant. This chapter will focus on those studies which deal with mechanisms of nickel-induced inhibition of plant metabolism that were observed under environmentally relevant conditions, preferably in vivo.

3.3.1.

Inhibition of Root Function

For terrestrial higher plants, the roots are usually the first target organ of any heavy-metal-induced damage, since they are in direct contact with the heavymetal-containing soil solution and almost all heavy metal uptake occurs through Met. Ions Life Sci. 2, 31–62 (2007)


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this organ. Inhibition of root functions has been observed in the environmentally relevant (e.g., for waste leachate, see above) concentration range. A 15% decrease of root growth was observed at 2.5 µM Ni2⫹ in a particularly sensitive strain of Silene paradoxa growing on nutrient solution in sand culture [102]. Gajewska and Sklodowska [114] found a significant decrease in root growth (fresh weight and length) starting at 10 µM Ni2⫹. Pandolfini et al. [115] measured root lipid peroxidation already at 20 µM Ni2⫹ in Triticum. Demchenko et al. [116] observed inhibition of root development only at 100 µM Ni2⫹, when cell elongation decreased and later cell division stopped. But 1 µM Ni2⫹ in the same experiment already led to subtle changes in the morphology of the rhizodermis, the relevance of which remained unclear. At the root level, nickel concentrations in the low micromolar range (measured with nutrient solution in sand culture) may also inhibit uptake of other essential trace elements such as copper and iron [117–120]. This may be due to competition for binding to, and transport by, metal transport proteins. Clearly, more research is required in this area, with environmentally relevant concentrations and more detailed studies on damage mechanisms.

3.3.2.

Inhibition of Photosynthesis

The photosynthetic apparatus, both its primary photochemical side and its biochemical carbon-fixing part, is one of the most important sites of inhibition by many heavy metals [4]. In all studies investigating this, a much stronger inhibition was found for photosystem II compared with photosystem I. The relative importance of specific inhibition sites, however, strongly depends on the nickel concentration, the irradiance conditions, the organism under investigation and the metal. Most of the work has been carried out with copper and cadmium as the most toxic metals; nickel has often been treated only as a side-aspect. Mechanisms of nickel-induced damage to photosynthesis include the formation of heavy-metal-substituted chlorophylls (hms-Chls) (reviewed by Küpper and Kroneck [4], nickel specifically investigated by Küpper et al. [98]). Substitution of Mg2⫹ in chlorophyll (Chl) by nickel results in an impairment of the correct function of the light-harvesting antenna because nickel-substituted chlorophylls (Ni-Chls) are not suitable for photosynthesis for many reasons as discussed in detail in a recent review by Küpper et al. [121]. The most important reasons are: (a) in contrast to Mg-Chl, Ni-Chls do not bind axial ligands [122], which are necessary for correct folding of the pigment–protein complexes [123,124]; (b) Ni-Chls have a very unstable singlet excited state that relaxes thermally [125], so that in solution they do not fluoresce and in vivo they do not transfer captured excitons towards the reaction center, but act as exciton quenchers [126]. In view of the latter fact, it is not surprising that only a few percent of the total Chl has to be converted to Ni-Chls for complete inhibition of photosynthesis [98] and that their exciton quenching leads to a strong decrease of fluorescence during shade Met. Ions Life Sci. 2, 31–62 (2007)

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reaction [98,127]. In metal-sensitive plants, lethal formation of Ni-Chls occurs in the micromolar concentration range, making it an environmentally relevant process [98]. However, studies on this process using sublethal nickel concentrations have not yet been carried out, since the authors used only copper and zinc in their later studies on hms-Chl formation. With higher nickel concentrations leading to high percentages of Ni-Chls in the plants, the strongly blue-shifted red absorbance maximum of Ni-Chls leads to a blue shift of the absorbance maximum of the extracts [98]. In most studies on nickel toxicity published so far, heavy metals were applied to Chlorophyta (higher plants and green algae) in high irradiance or without a dark phase. Under such conditions, direct damage to the PS II reaction center (PS II RC) occurs instead of the formation of Ni-Chls in the antenna. This reaction has been named the ‘sun reaction’ by Küpper et al. [98]. After inhibition of photosynthesis, under high irradiance the antenna pigments are degraded [98,128]. However, bleaching during sun reaction is slower than the Mg substitution during shade reaction, so that the very low ratio of Mg substitution during sun reaction cannot be caused by competition with the bleaching process [98]. Under some conditions, effects on photosynthesis differing from those discussed above (shade reaction, sun reaction) have been found. Using a long light period and an intermediate irradiance during severe nickel toxicity stress, Krupa et al. [129] observed a stronger inhibition of the light-adapted steady-state photosynthesis compared with the dark-adapted initial photosynthetic performance in Phaseolus vulgaris. Krupa et al. hypothesized that this effect indicates an inhibition of the Calvin cycle as the primary event, but it could as well be caused by a nickel-induced disturbance of the regulation of energy dissipation in the light reactions.

3.3.3.

Oxidative Stress

Many studies have dealt with oxidative stress under heavy metal stress, not only for redox-active metal ions such as Cu2⫹/Cu⫹, but also for metal ions that are usually not redox active physiologically, such as Ni2⫹. Lipid peroxidation was observed in Triticum roots, starting at 20 µM Ni2⫹ [115] and in algae already at 10 µM Ni2⫹ [130]. As mentioned already in Section 3.3.1, Gajewska and Sklodowska [114] found a significant decrease in root growth (fresh weight and length) in Pisum, starting at 10 µM Ni2⫹ applied for 9 days. This was correlated with an increased ascorbate peroxidase activity in the leaves, which can be interpreted as an indicator of oxidative stress. But surprisingly, the increase in ascorbate peroxidase occurred only in leaves, while in roots the activity of this enzyme decreased with increasing Ni2⫹ concentration. Similarly, glutathione-S-transferase activity was four times more induced in the leaves than in the roots. These observations may show that the Met. Ions Life Sci. 2, 31–62 (2007)


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Ni-induced oxidative stress resulted from malfunctioning photosynthesis, which certainly occurs only in the aboveground parts of the plant. The decrease of antioxidative enzymes in the roots could also be a direct effect of nickel, which in nonaccumulators is present in roots at much higher concentrations than in shoots and may inhibit uptake of essential trace elements (see Section 3.3.1.).

3.3.4.

Other Effects

In principle, heavy metals can have a mutagenic effect on the genome, but so far there is no convincing evidence that such effects play a significant role in in vivo heavy metal toxicity in plants. Manusadzianas et al. [97] reported a strong ATPase inhibition at sublethal levels of nickel: 90 µM IC50, in comparison to a lethal concentration (4-day LC50) of 290 µM. Nickel may induce a disturbance in the uptake of essential microelements as mentioned in Section 3.3.1. But while this was confirmed for roots by several authors [117-120], Quinn et al. [131] showed that it does not occur in the green alga Chlamydomonas. Instead, these authors found a nickel-induced upregulation of several copper-deficiency response genes. As a result, these genes remained active, even under conditions of excess copper. A break-up of colonies of the floating higher aquatic plant Lemna was observed already at 5 µM Ni2⫹ (or 0.2 µM Cu2⫹) within 6 h of exposure [101].

3.4. Mechanisms of Resistance Against Nickel Toxicity Plants have developed a number of strategies to resist the toxicity of heavy metals which has been extensively reviewed by many authors (e.g., Prasad and Hagemeyer [5], Küpper and Kroneck [4], Meharg [132]). As for mechanisms of damage, this section will focus on recent research and nickel-specific aspects. While metal tolerance is a constitutive feature in plants normally growing on metal-rich habitats such as hyperaccumulators (see Section 4), in Scenedesmus acutus the defense mechanisms were activated only at an elevated level when the cells were actually stressed by metal toxicity [130].

3.4.1.

Exclusion and Sequestration

A very common mechanism of nickel tolerance is exclusion. In Triticum, for example, Ni2⫹ toxicity triggered an enhanced activity of extracellular peroxidases that function in the production of phenolic compounds [115]. Such compounds, in particular lignin, serve to reduce the unspecific permeability of roots and thus reduce heavy metal uptake. However, an increase in tannins, a group of phenolics characteristic for conifers, was also found in needles of Scots pine exposed to elevated (already at 5 ppm) nickel in the soil [133]. Since in this case Met. Ions Life Sci. 2, 31–62 (2007)

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the tannins would not reduce Ni uptake, their increased production may be an unspecific stress response. Another exclusion mechanism may be mediated by ectomycorrhiza. It was found that nickel resistance in nickel-excluding species of pine, e.g., Picea glehnii, was correlated to a much increased (compared with nonresistant pine species) association of the roots with ectomycorrhizal fungi [120,134]. However, the physiological/biochemical mechanism of the suppression of Ni uptake in the plant by the association with these fungi remained unresolved. It is possible, for example, that the fungi excrete strong metal ligands that reduce nickel bioavailability. But at present it cannot be excluded that the correlation between increased mycorrhizal infection and increased Ni resistance was purely by chance. The latter impression is reinforced by the work of Ahonen-Jonnarth and Finlay [135], who found no reduction, but enhancement of nickel uptake after mycorrhizal colonization of pine (Pinus sylvestris) seedlings. It also cannot be excluded that the effect differs greatly depending on the particular species of tree and fungus. Clearly, more research is required in this area. Many plants detoxify heavy metals by sequestering them in the vacuoles [4]. This plant-specific metal detoxification strategy (animal and bacterial cells do not possess this organelle) provides an efficient form of protection because the vacuole does not contain any sensitive enzymes. When vacuolar sequestration is the major detoxification mechanism, nickel tolerance is often associated with elevated nickel accumulation; an extreme form of this sequestration is found in hyperaccumulator plants (see Section 4). In most heavy-metal-tolerant plants, the vacuolar sequestration occurs mainly in nonphotosynthetic cells of the epidermis, reducing toxicity to the heavy metal sensitive photosynthetic apparatus [95,136–139]. Both exclusion and vacuolar sequestration are metal transport processes against the concentration gradient, so that they require an active transport system. So far, knowledge about such transport systems is limited, in particular for Ni, as described in Section 2. Protein families involved in vacuolar sequestration may be the Nramps, CDFs, and CAXs (reviewed by Hall and Williams [141a]), but almost none of this research deals with Ni. Only one Ni-specific transporter for vacuolar sequestration has been characterized and it belongs to the CDF family. This is MTP1t2, closely related to MTP1t1, ZAT, and ZTP transporting mainly Zn [68,141b,141c].

3.4.2.

Binding by Strong Ligands

The main alternative to heavy metal tolerance by exclusion from the cell or sequestration inside the cell is binding to strong detoxifying ligands that prevents unwanted reactions of the metal ion with other ligands, for example proteins or chlorophyll, in the same physiological compartment. This has been studied for a long time and extensively reviewed [4,142,143], so that only aspects with particular importance for nickel will be discussed here. Met. Ions Life Sci. 2, 31–62 (2007)


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While phytochelatins are the most important cadmium chelators in nonaccumulator plants, they do not seem to play any role in nickel resistance. They have much lower affinity for nickel than for cadmium [144], so that is was not surprising that they were not found to be associated with nickel under Ni-induced stress in vivo [145]. Similarly, the main metal detoxifying ligands in animals, metallothioneins, do not seem to bind nickel in plants [143]. A group of ligands that are often associated with metals in plants are organic acids, such as citrate or malate (reviewed, e.g., by Rauser [142], Küpper and Kroneck [4]). Elevated nickel has been found to enhance the production of organic acids in various species. An extreme case of almost exclusive binding of nickel by organic acids is the latex of the Ni hyperaccumulating tree Sebertia acuminata [146] (see Section 4.2). Plants seem to use monomeric amino acids for binding nickel. This was initially observed in xylem sap, without identification of the specific amino acid, by Cataldo et al. [147]. Before this, the nonproteogenic amino acid nicotianamine was shown to bind nickel with high affinity, so it may be involved in nickel transport [73]. A very recent study by Douchkov et al. has shown that ectopic expression of nicotianamine synthase results in increased nickel tolerance in transgenic tobacco [148]. However, even this study does not prove that nicotianamine is involved in nickel metabolism or tolerance in normal plants because binding of nickel by nicotianamine under natural conditions has still never been shown. It is possible, for example, that under normal conditions there are better (stronger, easier to synthesize, easier to load/unload, etc.) ligands for nickel. In hyperaccumulators, histidine clearly has been shown (by X-ray absorption spectroscopy of xylem sap and tissue) to bind nickel in the xylem sap [149], as discussed in greater detail in Section 4.2. Also for histidine, recent studies have shown that enhanced synthesis leads to increased nickel tolerance (overexpression in Arabidopsis thaliana: Wycisk et al. [150], selection of Ni-resistant Nicotiana tabacum cell lines: Nakazawa et al. [151]). Another important chelator of metals, including nickel, are the cell walls, as far as the metal is transported apoplastically. While this is not the case in the shoots of hyperaccumulators [95], it is of importance in the roots of any plant because the cell walls inevitably come into contact with the soil solution [142]. Further, in water plants (including algae) the cell walls come into direct contact with the heavy-metal-containing medium and act as a cation exchanger.

3.4.3.

Other Mechanisms

The problems of oxidative stress are counteracted by the antioxidative defense system; upregulation of intracellular peroxidases was found in Triticum stressed with 20 µM Ni2⫹ [115]. Randhawa et al. [130] observed that Ni treatment induced an increased ratio of reduced:oxidized glutathione in a resistant strain of Met. Ions Life Sci. 2, 31–62 (2007)

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Scenedesmus. The opposite would have to be expected under Ni-induced oxidative stress, and indeed was observed in a sensitive Scenedesmus strain under the same conditions. This interesting observation indicates that, in the resistant strain, Ni stress induced a defense mechanism that ultimately overcompensated the glutathione oxidation. Enzyme assays in the same study showed that this defense mechanism was based on a much higher expression of glutathione reductase in the resistant strain. While the expression as such was constitutive, the enzyme was activated only in response to Ni2⫹ stress. Because of the sensitivity of the photosynthetic apparatus to heavy metal toxicity, plants have developed strategies that specifically protect this part of their metabolism, in addition to the general metal detoxification mechanisms described in Sections 3.4.1–3.4.3. In particular, in plants that accumulate large amounts of metals, the highest metal accumulation is usually found in cell types that are not photosynthetically active (see Sections 3.4.1. and 4.2.). Under conditions of heavy metal toxicity, hyperaccumulator plants seem to reduce the binding of Ni to essential sites (such as Chl) by accumulating the Mg that would be replaced by Ni [95] (see Section 4.2). In this context it is interesting to note that trees grown on high Ni in sand culture exhibited reduced Ni toxicity if an elevated level of magnesium was supplied simultaneously [120].

4.

NICKEL HYPERACCUMULATION

Plants have developed a number of strategies to resist the toxicity of heavy metals, as discussed in the previous section. Plants that actively prevent metal accumulation inside their cells are called excluders; these represent the majority of metal-resistant plants [6]. Other resistant plants deal with potentially toxic metals in just the opposite way, i.e., they actively take up metals and accumulate them. This phenomenon was discovered as early as 1885 by Baumann [152] for zinc accumulation in Thlaspi calaminare, later renamed to Thlaspi caerulescens ssp. calaminare [153]. These plants, which have been named ‘hyperaccumulators’ [7], are able to actively accumulate several percent of metals in the dry weight of their aboveground parts. This ability provides a promising approach for both cleaning anthropogenically contaminated soils (phytoremediation) and for commercial extraction (phytomining) of metals from naturally metal-rich (serpentine) soils (see Section 4.3.). By now more than 400 species of hyperaccumulators for diverse metals have been identified in about 50 plant families in many parts of the world, but representing less than 0.2% of all plant species [154,155]. Of all hyperaccumulators identified, about 75% are nickel hyperaccumulators. The best-known genus among these is Alyssum; a large number of species belonging to this genus accumulate nickel. Many other plant genera from various families hyperaccumulate nickel as well; two particularly noteworthy species among these are the tree Sebertia acuminata that accumulates up to 26% nickel in the Met. Ions Life Sci. 2, 31–62 (2007)


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dry weight of its latex [146], and the high-biomass herbaceous plants Alyssum bertolonii and especially Berkheya coddii [156–158] that are profitable ‘phytomines’ (see Section 4.2.).

4.1.

Biological Function of Nickel Hyperaccumulation

Many studies have shown that metal hyperaccumulation serves as a defense mechanism against herbivores and pathogens [159–163]. Herbivores that were given the choice between Senecio coronatus plants grown on different nickel levels chose those plants which accumulated the lowest amount of metal [161]. This effect seems to be directly caused by the metal accumulation, and not related to other chemical defenses in hyperaccumulators [164]. On the contrary, Davis and Boyd [164] have shown that hyperaccumulators have less need for other chemical defenses, so that they synthesize less allelochemicals than nonaccumulator species of the same genus. A very recent study has shown that intelligent herbivores such as locusts are not deterred from hyperaccumulators directly (e.g., by taste or appearance of the leaves or immediate toxic effects) but by a postingestive feedback mechanism involving associative learning [165]. The chemical defense of hyperaccumulators does not only work against herbivores, but also against bacterial [166] and fungal pathogens [166,167]. While it is clear from those studies that hyperaccumulation does protect against a broad range of herbivores and pathogens, like any other defense strategy it has limitations. The Ni hyperaccumulator Streptanthus polygaloides was more susceptible to turnip mosaic virus than the related nonaccumulator S. insignis, in particular when Ni-fertilized plants were compared [168]. A very recent study [163] testing the effect of nickel hyperaccumulation in Streptanthus polygaloides on different types of herbivores has shown that hyperaccumulation is an efficient protection against tissue-chewing herbivores, but much less efficient against herbivores feeding on vascular tissue. This is logical in view of the preferentially epidermal sequestration of the metal in most hyperaccumulators (see Section 4.2). In this context it should also be mentioned that hyperaccumulated metals may be spread from hyperaccumulators back into the environment via those herbivores that are not deterred by the high heavy metal content [169]. Since the nickel content accumulated in the plant can easily be controlled by the nickel concentration in the growth medium, hyperaccumulators may be an ideal model for a systematic study of plant-pathogen/herbivore interactions, as discussed by Pollard [170]. In addition to the protection against herbivores and pathogens, it was proposed that hyperaccumulation may serve as ‘elemental allelopathy’ [171]. This study suggested that hyperaccumulators increase the nickel concentration in the surface soil next to them and thereby inhibit the growth of nonaccumulators competing for space and nutrients. At the same time, the elevated metal concentrations would Met. Ions Life Sci. 2, 31–62 (2007)

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encourage growth of hyperaccumulator seedlings [95]. Elevated metal concentrations in the surface soil under hyperaccumulator plants were also measured by other authors [172,173]. Further support for this hypothesis was provided by a recent work of Zhang et al. [174] who investigated the degradation of Alyssum murale biomass in soil. Another alternative hypothesis about the biological role of hyperaccumulation is the increase of osmotic pressure for increased tolerance to the drought stress that often characterizes the natural habitats of hyperaccumulators. Results concerning this hypothesis are, however, contradictory. A study with the Ni hyperaccumulator Alyssum murale and the Cd/Zn hyperaccumulator Thlaspi caerulescens [175] showed that neither nickel in Alyssum murale nor zinc in Thlaspi caerulescens ameliorated growth inhibition by drought stress. And drought stress did not influence the accumulation of these metals. In contrast, in Stackhousia tryonii it was found [176] that the accumulation of nickel (measured in the dry weight of the shoots) increased with increasing drought stress, leading the authors to the conclusion that hyperaccumulation may have a role in protection against drought stress by osmotic adjustment. However, these authors did not make the reverse test as carried out by Whiting et al. [175], i.e., they did not test whether plants grown on high Ni really become more resistant towards drought stress compared with plants grown on low Ni. Another recent study [177] again questioned the role of hyperaccumulation as an osmotic adjustment. Nickel and zinc nutrition and accumulation did not influence in any way the salt tolerance of the nickel hyperaccumulator Alyssum murale or the zinc hyperaccumulator Thlaspi caerulescens. Clearly, further work is required in this area.

4.2.

Mechanisms of Nickel Hyperaccumulation

The mechanisms by which hyperaccumulator plants accumulate the enormous amounts of heavy metals in their shoots, and prevent phytotoxicity of these metals, have been a subject of many studies, in particular during the past decade, but many of these mechanisms are still under debate (reviewed, e.g., by Pollard et al. [178]). Work by Kerkeb and Krämer [179] on the Ni hyperaccumulator Alyssum lesbiacum has shown that the elevated histidine levels of some hyperaccumulators [149] play a role in xylem loading. Using X-ray absorption spectroscopy, Krämer et al. [149] did not find any sulfur ligands in the xylem sap of Ni hyperaccumulating Alyssum species, but found nickel binding by free histidine, which seems to be a general metal ligand in plants (see Section 3.4.2.). Histidine synthesis in nickel hyperaccumulators is much higher than in nonaccumulators, but it is constitutive and not nickel induced. The expression of the enzymes in its biosynthetic pathway is not changed by nickel nutrition [180,181]. Nickel in the hyperaccumulating tree Sebertia acuminata is bound mainly by citrate, as revealed by NMR Met. Ions Life Sci. 2, 31–62 (2007)


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spectroscopy of the latex of this tree [146]. Similar results were obtained for the complexation of nickel in roots of Alyssum bertolonii [182]. As for histidine, the synthesis of organic acids in hyperaccumulators seems to be constitutive and not metal induced. A number of recent studies have indicated that metal compartmentation may be a key mechanism of nickel tolerance in hyperaccumulator plants. Metals have been shown to be preferentially sequestered in subcellular compartments and cell types where they do the least damage to photosynthesis. Hyperaccumulators generally seem to sequester nickel in the vacuole [95,183,184], which is the ideal storage compartment for toxic substances since it is physiologically inactive. Most hyperaccumulator plants have been found to accumulate nickel in the epidermis [95,139,183–185], which makes sense for two reasons. First, the epidermis is the tissue ingested first by a tissue-chewing herbivore, so the epidermal storage is most efficient for anti-herbivore defence. Second, except for the stomatal guard cells the epidermis is not photosynthetic, so that storage in this tissue reduces the danger of nickel toxicity. A functional differentiation inside the epidermis, leading to accumulation mainly in large, non-photosynthetic storage cells and exclusion from the photosynthetic guard cells was found and proposed to further reduce toxicity [95,136–139]. The strong sequestration of metals into the vacuoles makes hyperaccumulators inefficient in using these metals as micronutrients; they need much higher concentrations of these metals for normal growth than all other plants [95]. Two noteworthy exceptions from the described pattern of metal compartmentation have been found. Old leaves of the Ni hyperaccumulator Berkheya coddii accumulate most of the metal as nickel silicates in the cell walls [186], and in the Ni-hyperaccumulating tree Sebertia acuminata the latex has by far the highest Ni concentration [146]. When metal concentrations in hyperaccumulators reach toxic levels, several additional resistance mechanisms may become activated. In the mesophyll of Nistressed Ni-hyperaccumulating Alyssum and Thlaspi species the concentrations of nickel rises faster than in the epidermis, suggesting that the epidermal sequestration mechanism becomes overloaded so that additional storage sites are needed [95,184]. But even under these circumstances the plants seem to avoid widespread nickel toxicity in the mesophyll, since the Ni accumulation occurs only in a small subset of the mesophyll cells, which are possibly sacrificed in an ‘emergency defense’ [95]. Under these stress conditions, an increase of Mg content was found in the same mesophyll cells, which was interpreted [95] as a defense against the replacement of Mg2⫹ in Chl by heavy metals [98]. Hyperaccumulators seem to be better equipped than normal plants to deal with nickel-induced oxidative stress. Although phytochelatins are not involved in nickel resistance (see Section 3.4.2.), Freeman et al. [187] found a much increased glutathione biosynthesis in various nickel-hyperaccumulating Thlaspi species. Furthermore, overexpression of a key enzyme in glutathione biosynthesis, serine acetyltransferase, increased nickel resistance in the non-accumulator Arabidopsis thaliana. Met. Ions Life Sci. 2, 31–62 (2007)

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4.3. Use of Nickel Hyperaccumulator Plants for Phytomining and Phytoremediation Many hyperaccumulators have a good potential to be used for phytoremediation, i.e., to extract and remove heavy metals from anthropogenically contaminated soils, which was first proposed by Chaney [188] and intensively investigated in the following years (recent reviews: [4,189–191]). Some of them even allow for commercially profitable phytomining, i.e., the extraction of metals (mainly nickel) from naturally heavy metal rich soils that are not directly usable as metal ores. After burning of the plants, their ash can be used as a metal ore (first proposed by Baker and Brooks [192]). Nickel was the first metal for which the economic feasibility of phytomining was shown and some nickel hyperaccumulators hyperaccumulate the even more valuable cobalt as well [158,193]. Nicks and Chambers [194] yielded a crop of nickel of equal value compared with an average crop of wheat by planting Streptanthus polygaloides on a metal-rich soil in California (USA). Furthermore, they showed that by burning these plants it is possible to yield, with low input of energy, a high-quality bio-ore (the plant ash) containing about 15% nickel. In another study, between 9 and 82% Ni were obtained in the bio-ore, depending on the plant, growth conditions, and the burning of the biomass [195]. Berkheya coddii has been known as a high-biomass (over 20 t/ha) Ni hyperaccumulator since the work of Anderson et al. [196]. Robinson et al. [156,158] carried out comprehensive studies of metal uptake, and showed that fertilization with sulfur and nitrogen greatly increased Ni and Co hyperaccumulation. Thus, their work has demonstrated that this species is a very promising candidate for both phytoremediation and phytomining. This has been confirmed by field trials in a recent study, which demonstrated that this species easily yields 110 kg nickel per hectare per year [197], and even 170 kg should be possible [198]. At current (February 2006) market prices of around 12.5 €/kg, that would be equivalent to around 2100 €, about four times the price of an average crop of wheat as calculated by Anderson et al. [199]. Problems in phytomining with B. coddii may occur when cobalt occurs in the soil. Although cobalt is hyperaccumulated as well, it is rather toxic for this species and reduced the Ni bioaccumulation coefficient from around 100 to 22 according to Keeling et al. [200]. As an alternative, also Alyssum bertolonii has already been shown to produce high enough nickel yields per hectare for phytomining [157,193,198], which now has already been put into commercial operation (reviewed by McGrath and Zhao [201] and Chaney et al. [190]) despite the doubts about the suitability of this species for phytomining expressed by Brooks et al. [197]. Phytoextraction, both for phytoremediation and for phytomining, depends on the pH of the soil. In contrast to most other metals, nickel uptake in hyperaccumulators increases with increasing pH [202]. Soil moisture, in contrast, does not seem to affect hyperaccumulation in Thlaspi, Alyssum, and Berkheya, so that



53

these hyperaccumulators may be applied in a wide range of climatic conditions [203]. A complete technological guideline for commercial phytomining was recently developed, based on field trials [204]. Several other large plants hyperaccumulate nickel, but their potential for phytoremediation and phytomining has not yet been tested. An extreme case seems to be the widespread tropical genus Phyllanthus (Euphorbiaceae). The Cuban species P. ⫻ pallidus was found to accumulate up to 6% nickel in its dry mass [205], and many others contained between 3 and 4% nickel; only the Australian Phyllanthus species are non-hyperaccumulating [206]. Phyllanthus species are medium sized (1–5 m high) shrubs; despite their size and extreme nickel contents (in particular in the Cuban species), no study has been published so far investigating the potential of Phyllanthus species for phytoremediation and phytomining as discussed by Reeves [206]. It has been repeatedly proposed to use non-accumulator crop plants for phytoremediation, because they may produce a higher biomass than hyperaccumulators [207]. This way of thinking is, however, misleading, as discussed in detail in the recent review of Chaney et al. [190]. First, non-accumulator crop plants such as maize or alfalfa are rather metal sensitive, so that on the contaminated soils to be remediated they would not produce a large biomass. Second, their bioaccumulation coefficient (metal concentration in plant divided by metal concentration in soil) is low and often even below 1, leading to a dilution rather than a concentration of the metal and thus to enhanced problems of disposal [208]. In contrast, hyperaccumulators strongly concentrate the metal with bioaccumulation coefficients up to 100 (discussed, e.g., by Keeling et al. [200], McGrath and Zhao [201], and Chaney et al. [190]).

5.

OUTLOOK

In nickel nutrition, it is most obvious that further research is needed in order to characterize the biochemical mechanisms of nickel uptake and translocation. For example, not a single one of the genes encoding (putative) nickel transporters has been isolated as a protein. As a result, the mechanisms of their function and their basis in the protein structure remain unknown. In the field of nickel toxicity, more effort is needed in revealing specific mechanisms of inhibition under controlled conditions that are a good simulation of the situation occurring in polluted environments. So far, in many cases either too high concentrations were applied in order to force effects to occur within a very short time, or poorly defined soil conditions were applied so that the influence of individual factors remained unclear. It is often argued that instead of using natural hyperaccumulators for phytoremediation and phytomining, genetically engineered plants should be used.


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Looking at the results of classical selection breeding of hyperaccumulators compared with attempts to create transgenic hyperaccumulators, the former approach appears much more promising, for the following reasons. Research on the mechanisms of hyperaccumulation as summarized in Section 4.2 has revealed that this process involves many different steps in diverse parts of the plant. Therefore, to re-create a hyperaccumulator by genetic engineering, one would have to modify the expression of many genes, in a tissue-specific way. This has not been achieved, not even in an approximation, in any study so far. Unless someone finds a general ‘switch gene’ that leads to the changed expression pattern of all the other genes involved in hyperaccumulation, transgenic plants that really accumulate as much metal as hyperaccumulators will remain science fiction. In contrast, field trials have shown that the biomass of hyperaccumulators can be dramatically increased by addition of fertilizer, natural selection, and classical breeding to reach levels that are economically attractive [190]. As a source for selecting species that are suitable for specific phytoextraction tasks, conservation of metallophyte biodiversity is of prime importance, as explained recently by a large group of internationally renowned scientists in the field [209].

ACKNOWLEDGMENTS This work was supported by the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie. Figure 2 was generously provided by Professor O. Einsle.

ABBREVIATIONS Chl EC50

EDTA GTP hms-Chl IC50

LC50 NiCoT

chlorophyll effective concentration 50% ⫽ the molar concentration of an agonist, which produces 50% of the maximum possible response for that agonist, measured in vivo ethylenediamine-N,N,N′,N′-tetraacetate guanosine 5′-triphosphate heavy-metal-substituted Chl, i.e., a chlorophyll where a heavy metal ion has replaced the natural Mg2⫹ as the central ion inhibitory concentration 50% ⫽ the concentration of an inhibitor required to achieve a 50% reduction of the in vitro activity of an enzyme (sometimes also used for the 50% activity reduction of a metabolic unit such as a chloroplast) lethal concentration 50%, meaning the concentration where 50% of all individuals die nickel cobalt transport protein



NiSOD PS II RC

55

nickel-dependent superoxide dismutase reaction center of photosystem II

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29. 30. 31. 32. 33.

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52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64.

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3 Nickel Ion Complexes of Amino Acids and Peptides Teresa Kowalik-Jankowska,1 Henryk Kozlowski,1 Etelka Farkas, 2 and Imre Sóvágó2 1

Faculty of Chemistry, University of Wroclaw, F. Joliot-Curie 14, PL-50383 Wroclaw, Poland

2

Department of Inorganic and Analytical Chemistry, University of Debrecen, H-4010 Debrecen, Hungary <[email protected]> <[email protected]>

1. INTRODUCTION 2. COMPLEXES OF AMINO ACIDS AND DERIVATIVES 2.1. Thermodynamic and Structural Studies of Nickel(II) Complexes of Amino Acids with Non-coordinating Side Chains 2.2. Thermodynamic and Structural Studies of Nickel(II) Complexes of Amino Acids with Side Chain Donors 2.3. Thermodynamic and Structural Studies of Nickel(II) Complexes of Amino Acid Derivatives 2.3.1. Complexes of Aminophosphonates 2.3.2. Complexes of Aminohydroxamates 2.3.3. Complexes of Oxime Derivatives of Amino Acids 3. COMPLEXES OF PEPTIDES AND RELATED LIGANDS 3.1. Nickel(II) Complexes of Peptides with Non-coordinating Side Chains

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3.2. Complexes of Peptides with Coordinating Side Chains 3.3. Nickel(II) Complexes of Peptides Containing Histidyl and Cysteinyl Residues 3.4. Complexes of Peptide Fragments from Histones, Protamines, and Other Biologically Important Proteins 3.5. Redox and/or Catalytic Reactions Involving Nickel(II) Complexes 3.6. Nickel(III) Peptide Complexes 4. FORMATION OF NICKEL(II) COMPLEXES UNDER BIOLOGICAL CONDITIONS: MODEL CALCULATIONS IN MULTICOMPONENT SYSTEMS 5. CONCLUSIONS ABBREVIATIONS REFERENCES

1.

77 80 82 86 91

94 96 97 98

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The basic properties of nickel and the coordination chemistry of nickel have been described in detail in Comprehensive Coordination Chemistry about 20 years ago [1,2]. Nickel(II) complexes of amino acids and peptides have also been part of a volume of the Metal Ions in Biological Systems series [3]. Ni(II), with amino acids and oligopeptides, forms complexes of two distinctly different stereochemistries: paramagnetic hexacoordinate and diamagnetic square planar species. Ni(III)-deprotonated peptide complexes can be easily obtained in solution by chemical or electrochemical oxidation of the corresponding Ni(II) complexes. The most stable electronic configuration of free Ni(II) is [Ar]3d8 which is also the ground state configuration in its complexes. In almost all of its six-coordinate complexes nickel(II) has a pseudooctahedral stereochemistry with a spin triplet as ground state (high-spin configuration). Three spin-allowed transitions are expected from the energy level diagram for d8 ions and the three observed bands in each spectrum may thus be assigned as 3A2g → 3T2g, 3A2g → 3T1g(F), 3A2g → 3 T1g(P). It is a characteristic feature of the spectra of octahedral Ni(II) complexes that molar absorbances of the bands are at the low end of the range (1–100) for octahedral complexes of the first transition series in general, namely, between 1 and 10. Magnetically, octahedral nickel(II) complexes have relatively simple properties. From both d-orbital splitting and energy level diagrams, it follows that Met. Ions Life Sci. 2, 63–108 (2007)

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they all should have two unpaired electrons, and this is always found to be the case, the magnetic moments ranging from 2.9 to 3.4 µB (BM), depending on the magnitude of the orbital contribution. For the vast majority of four-coordinate nickel(II) complexes, planar geometry is preferred. This is a natural consequence of the d8 configuration, since the planar ligand set causes one of the d orbitals (d x2y2) to be uniquely high in energy and the eight electrons can occupy the other four d orbitals but leave this strongly antibonding one vacant. Planar complexes of Ni(II) are thus invariably diamagnetic. They are frequently red, yellow, or brown owing to the presence of an absorption band of medium intensity (ε 60) in the range 450–600 nm, but other colors occur when additional absorption bands are present. In a regular square planar environment (D4h symmetry), three spin-allowed transitions corresponding to 1A1g → 1B2g, 1A1g → 1Eg and 1A1g → 1B1g are expected with low intensity (ε 50–200) due to the presence of the inversion center. The stabilization of high oxidation states requires ligands with high electron density and/or one or more negative charges in order to allow for some charge delocalization from the ligand to the metal. The ability to form strong σ bonds is a necessary requisite for a ligand to stabilize the 3 state in nickel complexes [4]. Nickel(III) complexes have been obtained with deprotonated peptides. Nickel(III) is usually observed in octahedral environments. Monomeric nickel(III) complexes are always paramagnetic at room temperature with effective magnetic moments ranging from 1.7 to 2.1 µB, corresponding to one unpaired electron. EPR spectroscopy has been extensively used to characterize nickel(III) complexes. The electronic configuration of nickel(III) is d7. In a tetragonal ligand field the ground configuration will be (d xzdyz) 4 (d xy)2 (dz2)1(d x2y2) 0 for a square planar or elongated octahedral complex and (d xzdyz) 4 (d xy)2 (d x2y2)1(dz2) 0 for a compressed octahedron. In the first case the EPR spectra will show g⊥ g 2.00 while in the latter situation the pattern g g⊥ 2.00 will be observed [5,6]. In strongly alkaline solutions, excess biuret (NH2-CO-NH-CO-NH2) with Cu(II) forms the well-known violet or biuret color which is the prototype for the complexation of four deprotonated amide nitrogens (from two biuret ligands). The reaction was described in 1848 [7], following up an earlier mention in 1833 that a corresponding reaction occurs with proteins, especially with egg albumin [8,9]. At the end of the 19th century Schiff concluded [8–10] from his systematic studies that the so-called biuret reaction, observed with biuret itself, also occurs with oxalamide (NH2-CO-CO-NH2) and with many of their derivatives. In addition, he noted that all those substances which exhibit the biuret color with Cu(II) show a yellow color with Ni(II) [8]. The correct structure for the alkaline violet biuret complex of Cu(II) was formulated in 1907 [11]. Investigations of ligand requirements for appearance of a biuret color continued [12] and included glycine peptides [13,14]. With glycinamide, triglycine, and higher peptides there is a cooperative transition from a green, paramagnetic, hexacoordinate complex to a yellow, diamagnetic, planar complex [15]. The acid–base properties of amino Met. Ions Life Sci. 2, 63–108 (2007)

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acids and peptides and their complexes have been described in a few reports [16–19]. It should be mentioned that deprotonated amides can be very effective in stabilizing nickel(III) as it was reported for biuret and substituted biurets [20].

2.

COMPLEXES OF AMINO ACIDS AND DERIVATIVES

Since the interest in Ni(II) complexes of amino acids arose a long time ago, a large amount of results for such systems have been published over the past decades. Also numerous excellent reviews, including some in the series of Metal Ions in Biological Systems (e.g., Vols. 2, 9, 23) or in the series of Specialist Periodical Report on Amino Acids, Peptides and Proteins issued by the Royal Society of Chemistry have appeared [3,16,21]. As a consequence, the main features, including stoichiometries, stabilities, structures, and spectral parameters of Ni(II)–amino acid complexes have already been clarified. Due to the exceptional importance of imidazole-N or mercapto-S as metal binding sites in biological sytems, Ni(II) complexes of both His and Cys have been extensively studied and discussed in detail in many reviews (see [16,21] and references therein). A great number of stability constants and thermodynamic values for various Ni(II)–amino acid complexes have been collected and also critically surveyed in special issues of Pure and Applied Chemistry [22] and in special data bases [23]. Therefore, only a brief discussion of the most important conclusions relating to Ni(II)–amino acid complexes is presented in this chapter. A list of the most common amino acids together with their corresponding acidity constants is shown in Table 1.

2.1. Thermodynamic and Structural Studies of Nickel(II) Complexes of Amino Acids with Non-coordinating Side Chains If the analytical concentrations are within the range of 103 101M, measurable interaction between these amino acids and nickel(II) starts around pH 5. Because at this pH the monoprotonated, zwitterionic form of the bidentate amino acids exists (HL), the first really occurring equilibrium process is the following competition reaction: Ni 2 HL → [ NiL] H The common bonding mode in the obtained complexes represents a bidentate coordination of the ligand via the carboxylate-O and amino-N donor atoms. In this way, a five-membered, thermodynamically stable chelate is formed with all the α-amino acidates, but a six-membered one with β -alaninate. If the pH is increased further, at appropriate nickel(II)-to-amino acid ratios, a second, then Met. Ions Life Sci. 2, 63–108 (2007)


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Table 1. Amino acids, H3NCH(R)COOH, and their acidity constants (pKa). pKa valuesa

Name

Symbol

R

α-Alanine Arginine

Ala Arg

CH3 + NH3

2.31, 9.70 2.03, 9.02 (12.1)

CH2(CH2)2NHC NH Asparagine Aspartic acid Cysteine 2,4-Diaminobutyric acid 2,3-Diaminopropionic acid Glutamine Glutamic acid Glycine Histidine

Asn Asp Cys Daba Dapa GIn Glu Gly His

CH2CONH2 CH2COOH CH2SH CH2CH2NH 3 CH2NH 3 CH2CH2CONH2 CH2CH2COOH H NH

CH2

2.15, 8.72 1.94, 3.70, 9.62 1.91, 8.16, 10.29 1.7, 8.19, 10.21 1.3, 6.68, 9.40 2.16, 9.96 2.18, 4.20, 9.59 2.36, 9.56 1.7, 6.02, 9.09

N Isoleucine Leucine Lysine Methionine Phenylalanine

Ile Leu Lys Met Phe

CH(CH3)(C2H5) CH2CH(CH3) 2 CH2 (CH2)3NH3 CH2CH2S(CH3)

2.21, 9.56 2.27, 9.28 2.19, 9.12, 10.68 2.10, 9.06 1.9, 10.41

CH2 Serine Threonine Tryptophan

Ser Thr Trp

CH2OH CH(OH)(CH3)

CH2 Tyrosine

2.17, 9.04, 10.11

Val β-Ala Pro

OH

CH(CH3) 2 H3NCH2CH2COOH +

N H2 a

NH

Tyr CH2

Valine β-Alanine Proline

2.13, 9.05 2.20, 8.96 2.34, 9.32

2.26, 9.49 3.53, 10.10 1.9, 10.41

CHCO2H

From [21–23].

a third, amino acid is also able to coordinate to the metal ion in stepwise processes. However, the hydrolysis of nickel(II) enters into competition with complex formation above pH 8 [3]. Evaluation of the stability constants and thermodynamic data of divalent 3d transition metal ions was made in numerous previous Met. Ions Life Sci. 2, 63–108 (2007)

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reviews and led to the conclusion that the sequence of data follows the Irving– Williams order [3,16]. Except for the Gly complexes, the logarithmic overall stability constants (log β) for [NiL] of simple bidentate α-amino acids fall into the region 5.0–5.5, for [NiL2] they are 10 and for [NiL3] close to 12.5–13. The reason for the somewhat higher stability of the Gly complexes has not been completely explained yet. Excellent correlation of logarithmic equilibrium constants of [NiL] complexes (except again for Gly and also the cyclic Pro) with the dissociation constant of the ammonium group of the free ligand (pKa) exists. Because of statistical reasons, the second stepwise stability constant (log KNiL2) is expected to be somewhat lower than log KNiL. Statistically, for octahedral complexes with bidentate ligands this difference is 0.7 log units. However, bulky chains, first of all because of steric reasons, might increase this difference somewhat, but on the other hand, non-covalent interactions, if they exist between the coordinated ligands, might decrease the difference. The latter is the case with Phe, where probably the interaction between the aromatic side chains of the two coordinating ligands favors somewhat the coordination of the second ligand. It is worth mentioning here that, since noncovalent interactions, in many respects, play a special role in biological systems, investigations of such interactions receive permanent interest. Ternary complexes (including those formed with Ni(II)) are often used as models in these studies and, not surprisingly, amino acids with bulky, aromatic or polar side chains are among the most preferred ligands [21,24–28]. With regard to the stereoselectivity in the formation of bis-complexes of amino acids with noncoordinating side chains, previous reviews clearly demonstrate that any differences are too small to be detected [3,21].

2.2.

Thermodynamic and Structural Studies of Nickel(II) Complexes of Amino Acids with Side Chain Donors

A large number of amino acids involve side chain donors which have the potential to form an additional bond to a metal ion, thus giving tridentate coordination. An obvious indication of tridentate coordination is a measurable increase in the stability constants relative to those of the truly bidentate amino acids. To quantify the extent of the side chain–metal ion interaction, a method was suggested by Martin [3] and also used in another review [16]. Based on this evaluation, no measurable intramolecular interaction is observed for nickel(II) and the guanidino side chain of Arg, or the ε-amino group of Lys (which would be able to form only an eight-membered chelate), or the phenolic group of Tyr (where the coordination of the side chain donor is sterically not favored), or the γ -amide O or N donor atoms of Gln. The weak basicity of the indole group does not provide a real possibility for its coordination, hence, Trp is expected to coordinate in a ‘normal’ bidentate way Met. Ions Life Sci. 2, 63–108 (2007)


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via the α-amino-N and carboxylate-O atoms, but the result calculated in chapter III of [16] indicates some interaction. Deprotonation of the very weakly acidic alcoholic group of Ser or Thr (pKa 14) is not induced by nickel(II), but a weak interaction of the non-deprotonated alcoholic group is suggested both in the solid state and in solution. As mentioned above, the γ -amide of Gln does not play any measurable role in the interaction with nickel(II), but a weak interaction via the somewhat stronger γ -carboxylate donor of Glu and a well measurable interaction via the much stronger γ -amino-N donor of Orn occurs [29]. A six-membered chelate can be formed by coordination of the side chain amide function of Asn. However, because of the poor donor properties of a non-deprotonated amide group the stability increase of [NiL] and [NiL2] complexes is only 0.4–0.5 log units per chelate. This weak coordination is the reason why a third coordinating Asn is able to displace the amide groups and the species [NiL3] , with a glycine-type coordination of the three ligands, is formed [30]. The rest of the amino acids contains strong side chain donors in suitable positions to form five- or six-membered chelates joined to the glycinate-type fivemembered one. Together with the amino-N, a six-membered chelate can be formed by the coordination of the β -carboxylate of Asp, or β -amino group of Daba, or imidazole-N(3) of His, while five-membered chelates form by coordination of the α-amino group of Dapa or the thiol group of Cys. Alternatively, the side chain donor may coordinate together with the α-carboxylate. Completely protonated forms of these ligands are able to release three dissociable protons in the measurable pH-range. The β -carboxylic acid group of Asp is quite acidic, but all the other side chain functions start to deprotonate only above pH 5 (His) or even higher (Table 1). As a consequence, the formation of protonated complexes is hardly important in the Ni(II)-Asp system, but with the other ligands, Daba, Dapa, His, and Cys it is important. Obviously, the monoprotonated ligands coordinate in a bidentate manner. In the coordination of the nonprotonated form of these ligands the side chain donor plays a crucial role and a tridentate coordination via two joined chelates is expected. Although there is a strong tendency for tridentate coordination with all of these ligands, differences between their nickel(II)-binding ability can also be found. For example, tridentate chelation allows the coordination of two ligands at most to a nickel(II) ion in an octahedral complex. However, this is the case only with Asp and His; with Daba, Dapa, and Cys, the situation is more complicated. Although the formation of bis-complexes with tridentate coordination of two Daba or Dapa is favored, some 1:3 complex also appears at high ligand excess [29]. Moreover, with Dapa [29] and Cys [21], under certain conditions, even a change in the geometry from octahedral to planar occurs. Cysteine does not coordinate nickel(II) in a tridentate way at all, but (S,N)-chelation occurs and a diamagnetic planar 1:2 complex results. Sulfur-bridged [Ni3L4] species are also suggested in this system [16,21]. Met. Ions Life Sci. 2, 63–108 (2007)

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To compare the stability of the different types of chelates with each other, pH-dependent conditional constants were calculated for mono-chelated nickel(II) complexes of some selected models, such as α-Ala and β -Ala (models for five- and six-membered (N,O)-type amino-carboxylate chelates), 1,2-diaminoethane (en) and 1,3-diaminopropane (pn) (models for five- and six-membered (Namino,Namino)chelates, respectively), and histamine (Hm) (a model for a six-membered (Namino,Nim)-chelate). The conditional stability constant can be defined as: n   α H [ L]  1 ∑ β Hi L [ H i ]  i1  

K ′ βNiL / α H

where βNiL are the corresponding overall stability constants taken from [21, 31, 32], [L] is the totally deprotonated form of α-Ala, β -Ala, en, pn, and Hm; n is the number of protons competing with the formation of the chelate (this value is 2 in all of these models), and βHiL is the corresponding protonation constant taken from Table 1 and [33]. The calculated logarithmic K values for the different chelates with Ni(II) as a function of pH are shown in Figure 1. It can be seen in Figure 1 that although the five-membered glycine-type chelate (line 1) is somewhat preferred to the others (it has the highest log K value) in the acidic region, there is no measurable formation of nickel(II) complexes in this pH

8 2 6 4 3

logK′ NiL

2

5

0 1 –2 –4 4

–6 –8 –10 2

3

4

5

6

7

8

9

10

pH

Figure 1. Conditional stability constants (log K) calculated for monochelated (1:1) Ni(II) complexes of α-Ala (5-N,O chelate, line 1), en (5-N,N chelate, line 2), Hm (6-Namino, Nim chelate, line 3), pn (6-N,N chelate, line 4), and β -Ala (6-N,O chelate, line 5) in dependence on pH. Met. Ions Life Sci. 2, 63–108 (2007)


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range (see the low numerical values of log K). As expected, the six-membered amino-carboxylate chelate (line 5) is always less stable compared with the fivemembered one. There is, however, a change in the order of log K values for the Hm-, en-, and pn-containing 1:1 complexes at about pH 5, 5.5 and 9 (lines 3, 2, and 4), respectively. As a consequence, above pH 5 His behaves more like a Hm derivative than an α-amino acid derivative. Above pH 5.5 the en-type character of Dapa becomes more and more dominant, but the stability of the six-membered pn-type chelate is comparable to that of the five-membered amino-carboxylate one only above pH 8 with Daba. A stable tridentate coordination mode of Dapa and especially Daba in their [NiL] complexes was indicated by the reduction potentials observed at dropping mercury electrode. From the redox potential of [Ni(His)], a π-acceptor behavior of the imidazole ring was also suggested [34]. The question of stereoselectivity in nickel(II) complexes of tridentate amino acids has been discussed in previous reviews [3,16,21]. While no stereoselectivity was found with potentially tridentate amino acids, such as Asn, Asp, Gln, Glu, a marked stereoselectivity was observed with His.

2.3. Thermodynamic and Structural Studies of Nickel(II) Complexes of Amino Acid Derivatives There is no doubt that amino acids are versatile starting materials, often used for the synthesis of different new ligands. The obtained derivatives show in many cases higher metal binding ability or special metal ion selectivity, and also biological recognition of such compounds is often quite effective because they incorporate amino acid(s). However, because the character and also the nickel(II)-binding ability of many derivatives of amino acids, such as Schiff bases [35–38], functionalized triazacyclononane [39], or diazacyclooctane derivatives [40], special thiourea derivatives of α-amino acids [41], N,N,N,N,tetrakis(carboxymethyl)-L-Orn [42], bis(picolyl)amine derivatives of Gly ethyl ester [43], or a new pentadentate tripodal ligand involving a glycinamide part [44], are far from the parent amino acids, these results are not discussed in this chapter. We also do not concentrate on derivatives for which only a few potentiometric results have been published (e.g., [45,46]). Results for the metal binding ability of bis(imidazolyl) derivatives of Gly, Phe, and His have been published in some recent papers [47–50]. In the next sections we focus on nickel(II) complexes of some amino acid derivatives in which either the carboxylic acid or the amino sites are modified. Namely, the nickel(II)-binding abilities of aminophosphonates (in these derivatives the carboxylate group is replaced by a phosphonate moiety, PO32), aminohydroxamates (where the carboxylate group is replaced by a hydroxamate group, CONHO), and oxime analogues (in which the amino group is replaced by an oxime moiety, N(OH)) are summarized below. Met. Ions Life Sci. 2, 63–108 (2007)

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2.3.1.

KOZLOWSKI et al.

Complexes of Aminophosphonates

Since versatile and useful biological activities can be attributed to aminophosphonates [51], there has been a special interest in the study of these compounds. Moreover, because some of the biological effects most probably correlate to the metal binding ability of these compounds, their metal complexes have been extensively studied, especially by Kozlowski’s group in Poland and also by the group of Kiss in Hungary; the results obtained are summarized in several reviews [21,52]. Because the carboxylate group of the parent amino acid molecule is exchanged for a phosphonate group, the effects caused by this change originate, first of all, from the differences of the two groups, such as their size, shape (flat carboxylic versus tetrahedral phosphonic), basicity, and charge [52]. Although predominantly Cu(II) complexes of aminophosphonic acids have been investigated, some results with Ni(II) have also been published [53–56]. Based on potentiometric and UV-Vis results, it has been established that with derivatives of simple aliphatic αamino acids the bidentate binding mode via amino-N and phosphonic-O atoms is predominant and in the maximum three chelates are able to coordinate to a Ni(II) ion in octahedral complexes. However, under acidic conditions, the more basic character of the PO2 3 group leads to the formation of protonated 1:1 complexes ([NiLH]) in which monodentate coordination of the phosphonate group can be assumed, but a chelating isomer (via coordination of NH2 and PO2 (OH)) cannot be excluded [52]. With the γ -phosphonic derivative of Glu, similarly to the parent amino acid, only bidentate coordination (via the α-amino-N and carboxylate-O) occurs, and the terminal phosphonate part cannot coordinate due to its distance from the five-membered chelate. However, the bonding mode with the derivatives of Asp (no matter which of the two carboxylate groups is exchanged for the phosphonate one) is tridentate, which allows the coordination of two ligands in the maximum in stepwise processes. The coordination of the second and especially the third ligand (only with the Glu derivative) is less favored for electrostatic reasons [54].

2.3.2.

Complexes of Aminohydroxamates

The hydroxamic acid group (which is a weak monoprotonic group with a pKa of about 8–9) is known as one of the most efficient metal ion-binding sites in natural systems [57]. Since both the natural and synthetic hydroxamic acids show numerous biological effects, there has been continuous interest in studies of such compounds. Natural hydroxamate-based compounds, siderophores, play a crucial role in microbial iron(III) uptake, storage, and transport [58], but the hydroxamic acids are also efficient chelating agents for many other metal ions and are effective inhibitors of metalloenzymes (e.g., nickel(II)-containing urease) [59]. Met. Ions Life Sci. 2, 63–108 (2007)


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Exchange of the carboxylate group of amino acids for a hydroxamate one results in the formation of aminohydroxamic acids. Simple aminohydroxamic acids can liberate two protons in overlapping processes in the pH range 5–10, one from the protonated amino group and one from the hydroxamic acid group. As a consequence, only the dissociation (protonation) microconstants can characterize the acid–base properties of the individual groups of the molecules. As it is known from previously determined microconstants, -NH3 is more acidic than the hydroxamic group in the case of α-alaninehydroxamic acid, but the trend is the opposite in β -alaninehydroxamic acid [60]. Although the -NH2 (CH3) group is also in α position compared with the hydroxamic one in sarcosinehydroxamic acid, the effect of the methyl substituent changes the acidity trend compared with α-alaninehydroxamic acid [61]. Due to the presence of the amino-N in chelatable position to the hydroxamate moiety, the aminohydroxamic acids are able to form not only the well-known hydroxamate-type five-membered (O,O)-chelate (Scheme 1, structure I), but also the (N,N) one (structure II) and, if the stability of the two chelates is comparable, the mixed type binding mode (structure III) is also possible. With Ni(II), the trans (N,N) coordination of two glycinehydroxamates in a square planar complex, trans-[NiL2], was proven by X-ray in the early 1980s by Brown et al. [62]. It was also established that one of the coordinated ligands releases its -NOH proton at high pH, which results in the formation of the even more stable planar cis-[NiL2H1] [63]. Many spectroscopic and pH-metric results obtained for nickel(II) complexes of simple aminohydroxamic acids in solution support the notion that the bis-complex, with a 4N coordination and planar geometry, which was found in the solid state, is especially favored [57]. Measurable complex formation starts at about pH 5.0–5.5 and practically at this pH the color of the samples turns yellow, indicating the formation of the planar bis-complexes. (Two characteristic bands, one in the range of 425–435 nm and another one in the range of 490–500 nm, assigned to 1A1g → 1A2g and 1A1g → 1B1g transitions, respectively,

Scheme 1. Met. Ions Life Sci. 2, 63–108 (2007)

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KOZLOWSKI et al. R

O

H2N

N

O H

Ni H2N

N

R

O

O

Scheme 2.

can be observed in the electronic absorption spectra.) Above pH 9, a new species, [NiL2H1] is formed, for which the binding mode shown in Scheme 2 is strongly suggested. The additional donor atom in the side chain of aminohydroxamic acids increases the number of possible coordination modes, but in most instances (derivatives of Phe, Tyr, DOPA, Orn, Lys, Arg, Met [64–66]) the above discussed coordination mode remains [57]. Significant effects of the side chain donors on the coordination mode were found only with derivatives of His [67], 2,4-diaminobutyric acid, and 2,3-diaminopropionic acid [65]. In the nickel(II)-histidinehydroxamic acid system, the complexes formed below pH 9 are octahedral or pseudooctahedral. Amino-N and imidazole-N chelated mono- and bis-complexes are suggested to form at lower pH, while following the deprotonation of the hydroxamic moieties, tridentate coordination (via amino-, imidazole-, hydroxamate-N) of the two ligands occurs leading to the formation of the planar [NiL2H1] complex (Scheme 2) [67]. Tridentate (NH2, NH2, Nhydrox) coordination also dominates for the derivatives of 2,4-diaminobutyric and especially 2,3-diaminopropionic acid. With the latter ligand the octahedral [NiL2] species, in which each of the two ligands is coordinated via two joined five-membered chelates, has a very high stability and the comparison of the nickel(II)- and iron(III)-binding ability of diaminohydroxamates, demonstrates nickel(II) preference for 2,3-diaminopropionohydroxamic acid above pH 6 [65]. Surprisingly, even the NH2 and the hydroxamic acid groups are in the β -position with respect to each other in β -alaninehydroxamic acid, the six-membered (N,N)-chelate is somewhat favored over the five-membered (O,O)-hydroxamate one and the planar 4N-coordinated bis-complex is formed with nickel(II) [57]. In addition to the 6-membered (N,N)-chelate, also the αcarboxylate takes part in the coordination of aspartic acid-β -hydroxamic acid [68] and octahedral complexes result. Tridentate coordination is also strongly supported by the results obtained with glutamic acid-γ -hydroxamic acid. In this latter case, complexes with different binding modes existing in equilibrium were suggested [69]. Met. Ions Life Sci. 2, 63–108 (2007)


2.3.3.

75

Complexes of Oxime Derivatives of Amino Acids

In the oxime derivatives of amino acids the amino group, which is usually an anchoring site for metal ions, is modified; hence, this modification is expected to cause a significant change in the coordination ability of the new ligand. According to the results obtained for nickel(II) complexes of several derivatives [70–73], these ligands are very efficient chelating agents of nickel(II). 2-(Hydroxyimino)propanoic acid (which is the oxime derivative of α-alanine) and its amide, as well as 2-cyano-2-(hydroxyimino)acetic acid and its amide, and also its ethane-1,2-diamine derivatives have been studied (Scheme 3). The alanine derivative coordinates to nickel(II) via its imino-N and carboxylate-O and the formation of bis-complexes with different protonation at oxime groups is especially favored. The deprotonation of the oxime hydroxyl group of one of the two coordinated ligands occurs at much lower pH compared with the free ligand. This is assumed to cause a strong shift of the electron density towards the coordinated imino nitrogen, which makes this donor more efficient in metal ion coordination. The stability constant of the octahedral [NiL2] 2 species is more than three orders of magnitude higher than that of the corresponding complex of the parent molecule, α-alanine, and only slightly weaker than that of the square planar [NiL2] complex of the very effective ligand, α-alaninehydroxamate (see above) [70]. In contrast to the alanine derivative, its amide forms a square planar bis-complex. In the complex [NiL2H1] , which is predominant at pH 9, the two imino-N and two deprotonated amide-N atoms are coordinated to the metal ion, and the cis arrangement of the two ligands is stabilized by the hydrogen


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bond existing between the deprotonated and nondeprotonated hydroxyl groups. Due to the strong electron withdrawing effect of the cyano group, the oxime OH becomes several orders of magnitude more acidic by exchanging the methyl for a cyano substituent in the alanine derivatives. As a result of this effect, the 2-cyano2-(hydroxyimino)acetate derivatives are somewhat less effective chelating agents for nickel(II) than the 2-(hydroxyimino)propanoate derivatives [71,72].

3.

COMPLEXES OF PEPTIDES AND RELATED LIGANDS

The systematic studies on the nickel(II) complexes of peptides go back to the early 1960s when Martin and coworkers reported that nickel(II) ions, similarly to copper(II), are able to induce deprotonation and metal ion coordination of the peptide amide functions [15]. The most important results obtained on the thermodynamic and structural properties of nickel(II) complexes of oligoglycines and their C-terminally amidated derivatives have already been reviewed by several authors [16–18,21]. It is clear from these compilations that there are some major differences in the complex formation processes of copper(II) and nickel(II) with peptide ligands. Copper(II) generally forms the common tetragonally distorted octahedral complexes with any oligopeptide ligand and deprotonation and metal ion coordination of amide functions take place in well-separated consecutive reactions with increasing pH. However, in the case of nickel(II) the coordination geometry of the complexes of dipeptides and the longer counterparts are completely different. Diglycine forms the pale blue, octahedral, paramagnetic nickel(II) complexes with [NiH2L2] 2 stoichiometry, in which the terminal amino, deprotonated amide, and carboxylate functions are the metal binding sites [74]. The deprotonation takes place in the alkaline pH range and generally high excess of ligand is required to avoid precipitation of nickel(II) hydroxide in basic solution. In contrast to diglycine, the interaction of nickel(II) and triglycine or larger peptides results in the formation of yellow, square planar, diamagnetic species [75] and the deprotonation of the two or three amide functions is highly cooperative. The origin of this cooperativity and the other principal properties of the nickel(II) complexes of the most common peptide ligands have already been reviewed [3]; hence, in this chapter we give an overview of only the most important or most recent results obtained on the role of various side chain residues in complex formation and also on their biological consequences.

3.1. Nickel(II) Complexes of Peptides with Non-coordinating Side Chains The above-mentioned differences in the coordination geometries of di- and tri(or higher) peptides also hold for the peptides containing the common noncoordinating side chains residues (e.g., the peptides of Ala, Leu, Val, Phe or Trp) Met. Ions Life Sci. 2, 63–108 (2007)


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[76–79]. Some more recent results, however, suggests that the insertion of bulky, non-natural residues into the peptide backbone can modify the coordination geometry of nickel(II)-peptide complexes. For example, the dipeptides of α-aminoisobutyric acid (Aib) formed low-spin, six-coordinated complexes, [Ni(H1Aib2)2]2, with nickel(II) which can be easily oxidized to a relatively stable nickel(III) species [80]. α,β-Dehydro amino acids (or ∆-amino acids) contain a double bond between the α- and β-carbon atoms and their insertion into the peptide sequence also has a significant impact on the coordination mode in peptide complexes [81,82]. The stoichiometry of the nickel(II) complexes of dipeptides containing C-terminal ∆Xaa residues were similar to those of the common dipeptides, but the formation of octahedral and square planar complexes was suggested for the [NiH2L2]2 species of Xaa-∆-Ala and Gly-∆-Ala, respectively [81]. The formation of only square planar nickel(II) complexes was detected if ∆-Phe was inserted into a tri- or tetrapeptide and the (Z)-isomer was the most effective one in the stabilization of the complexes formed [82]. The insertion of β -alanine into the peptide sequence changes the number of atoms within the chelates from five to six in the peptide complexes. In the case of nickel(II) this change does not affect the speciation and the basic coordination modes, but the overall stability of the complexes of dipeptides with (NH2, N, COO) and tripeptides with (NH2, N, N, COO) coordination modes follows the stability order: (5,5) (5,6) (6,5) (6,6) and (5,5,6) (5,6,5) (5,5,5) (6,5,5) (5,6,6) (6,5,6) (6,6,6), respectively [83].

3.2. Complexes of Peptides with Coordinating Side Chains It has been demonstrated in Section 2 that oxygen, nitrogen and sulfur donor atoms in the side chains of amino acids can significantly influence the metalbinding affinity of the ligands. Among them imidazolyl nitrogen donor atoms of histidyl and thiol sulfur atoms of cysteinyl residues are the most common and most efficient metal-binding sites and their complex formation reactions are discussed in Sections 3.3 and 3.4. In this section we focus on the complex formation of peptides containing alcoholic, phenolate or carboxylate oxygen, lysyl amino nitrogen and thioether or disulfide sulfur atoms in the side chains of natural amino acids. The results obtained on the nickel(II) complexes of various derivatives of peptides including the conjugates of some chelating agents will be discussed as well. Oxygen donor atoms of seryl, threonyl or tyrosyl residues generally can contribute to the overall stability of peptide complexes of transition metal ions, including copper(II) and zinc(II) and especially the hard trivalent metal ions. However, in the case of nickel(II) there is no unambiguous proof for the existence of Ni–O(alcoholic) or Ni–O(phenolate) binding in peptide complexes. On the contrary, the interaction of the aromatic side chains of phenylalanine Met. Ions Life Sci. 2, 63–108 (2007)

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or tyrosine with nickel(II) ion was suggested on the basis of NMR measurements. This type of stacking interaction was especially favored in the square planar nickel(II) complexes of tri- and tetra-peptides containing Phe and/or Tyr residues [84]. The β - and γ -carboxylate groups of aspartyl and glutamyl residues, respectively, are also potential metal-binding sites in peptide complexes. The stability enhancement of the β -carboxylate group of the aspartyl residue is generally more evident [85] because the terminal amino nitrogen and carboxylate oxygen atoms can form a six-membered chelate in the case of aspartic acid, while it is a seven-membered one for glutamic acid. The stability increase of various species can be observed both in copper(II) and nickel(II) complexes with the former being more pronounced, supporting that nickel(II) has a relatively low affinity to bind oxygen donor ligands [86,87]. Some peptides discussed in the previous sections contained also lysine residues, but the involvement of the ε-amino group of lysine in metal binding was not suggested in any of these cases [87–89]. The ε-amino group, however, can be involved in amide binding too, and this modification of the peptide backbone generally significantly increases the variety of complex formation processes. The nickel(II) binding affinities of γ -Glu-ε-Lys and Glu-ε-Lys were compared in a combined potentiometric and spectroscopic study [90]. In the former ligand the α-amino acid-like binding sites are well separated from each other and also from the amide function. As a consequence, the peptide amide deprotonation and coordination is hindered and a very stable [NiL] species is formed containing the amino acid binding sites supported by a macrochelate or loop structure. The formation of a similar species with outstanding thermodynamic stability was also suggested for the nickel(II)-Gly-Lys(Gly) system, indicating that nickel(II) has a high affinity to form macrochelate structures [91]. The ligands Glu-ε-Lys [90] and Asp-ε-Lys [91] also contain the ε-amino group in the amide bond, but it is in the chelating position with the terminal amino group which results in the exclusive formation of various dinuclear complexes. The sulfur atom is one of the most common metal-binding sites in proteins and it can be found in different chemical forms including thiols, disulfides, and thioethers. Among these, thiols have the highest affinity towards transition elements, including nickel(II), but this topic is discussed in the next section. The oxidation of thiols to disulfides, however, dramatically reduces the affinity of the sulfur atoms towards nickel(II). Although the existence of Ni–S(disulfide) bonds has been proven in the solid state for nickel(II) complexes of several model compounds [92], similar interactions were ruled out in the nickel(II) complexes of oxidized glutathione, (CysGly)2 and (GlyCys)2 [93,94]. The presence of the separated (NH2, COO) or (NH2, CO) binding sites, however, results in the formation of [NiL] complexes as the major species and their structure is stabilized by the formation of macrochelates. The studies reported on the nickel(II) complexes of the peptide hormones oxytocin and vasopressin came to similar conclusions supporting that Met. Ions Life Sci. 2, 63–108 (2007)


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the common oxygen and nitrogen donors of peptides in chelating positions are more efficient binding sites for nickel(II) than the disulfide sulfur atoms [95,96]. The formation of Ni(II)–S(thioether) bonds was suggested in the complexes of some nonproteinogenic derivatives of amino acids [97], but the systematic studies on the nickel(II) complexes of peptides of methionine revealed the existence of only very weak Ni(II)–S interactions in solution [98,99]. The similarity in the complex formation processes of triglycine and the tripeptides of methionine is especially remarkable in the square planar, diamagnetic (NH2, N, N, COO)-coordinated species, while a weak interaction of the thioether residue was suggested in the octahedral [NiL] complexes containing N-terminal methionyl residues. The various derivatives of peptides and their metal complexes have received increasing attention in the last decades. These molecules include the insertion of additional functional groups into the side chains of peptides or the replacement of amide and/or carboxylate functions with thioamides and phosphonic/phosphinic groups, respectively. The most interesting ligands are, however, the so-called peptide conjugates in which the peptide molecules are covalently linked to other efficient ligands including chelating agents and macrocycles. α-Hydroxymethylserine (HmS) is a nonproteinogenic amino acid containing an extra CH2–OH side chain on the α-carbon atom of serine and it can be found in several antibiotics. It has been reported that the insertion of HmS residues into di- or tripeptides, HmS-His and HmS-Hms-His, significantly increases both the copper(II) and nickel(II) binding capacity of these ligands compared with those of GlyHis or GlyGlyHis [100,101]. (Aminoalkyl)phosphonic and (aminoalkyl)phosphinic acids are the phosphorous analogs of the naturally occurring amino acids and their insertion into the peptide backbone provides an efficient way for the modification of the acid-base and complexing properties of the oxygen donor atoms of peptide ligands. Most of the publications, however, came to the conclusion that the carboxylate/phosphonate(phosphinate) substitution does not significantly influence the speciation and thermodynamic stability of the corresponding nickel(II) complexes [102–105]. In contrast with the low nickel(II) binding affinity of thioether and disulfide functions the insertion of the thiocarbonyl groups in the oligopeptide molecules critically changes the coordination ability of the peptide ligands. The Ni(II)– S(thioamide) bonded species of both di- and tetrapeptides predominate around the physiological pH, but this binding mode cannot prevent the deprotonation and coordination of amide functions in alkaline solutions [106–108]. In the last decade an increasing number of studies have been performed on the metal complexes of functionalized peptide molecules containing chelating agents or other biologically important ligands in the side chains. One group of these ligands includes the peptide derivatives of the chelating agent bis(imidazol2-yl)methylamine (BIMA) which can be linked to the C-termini of peptides via an amide bond. The results obtained on the amino acid derivatives of BIMA Met. Ions Life Sci. 2, 63–108 (2007)

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have already been reviewed [47–50] and discussed in the previous section. The data reported on the dipeptide-BIMA conjugates revealed that the presence of the bis(imidazolyl) chelating agent significantly enhances the nickel(II)-binding affinity of the peptides, especially below and in the physiological pH range, but similarly to the common peptides the amide functions will be the major metal ion binding sites in basic solutions [109]. Peptide nucleic acids (PNA) are synthetic analogs of DNA in which the natural phosphate–deoxyribose backbone is replaced by a peptide chain. It was found that the insertion of nucleobases, especially of thymine, has a critical impact on both the thermodynamic stability of nickel(II) complexes and the conformation of the ligands [110,111].

3.3. Nickel(II) Complexes of Peptides Containing Histidyl and Cysteinyl Residues Imidazole-N donor atoms of histidine and thiol sulfur atoms of cysteine are the most common metal-binding sites in proteins. The thermodynamic and structural properties of the copper(II) and nickel(II) complexes of peptides containing histidyl residues have been reviewed recently [18]. It is clear from these studies that the presence of histidine in any location of the peptide chain enhances the nickel(II)-binding affinity of the molecules. In the case of N-terminal histidyl peptides the stability enhancement is connected to the ‘histamine-like’ (NH2, Nim) coordination mode, while in the case of Xaa-His-Xaa and Xaa-Xaa-His-Xaa type peptides the outstanding stability of the (NH2, N, Nim) and (NH2, N, N, Nim) coordination modes, respectively, are responsible for the high nickel(II)binding capacity. The latter binding mode represents the simplest model of the metal-binding motif of albumin and its chemical and biological consequences are discussed in the forthcoming paragraphs. The nickel(II) complexes of a series of tetrapeptides have been studied to obtain information on the role of histidyl residues located in the fourth position from the N-termini. The imidazole-N donor atoms of histidyl residues were the primary metal-binding sites of the peptides in all cases, resulting in the formation of thermodynamically more stable species than those of the common tetrapeptides [112]. The nickel(II) complexes of peptides containing two and more His residues have also been studied in the last few years. The results of CD spectroscopic measurements revealed that protonation and metal ion coordination can induce folding of the linear octapeptide His(GlyHis)3Gly. The folding effect of copper(II), nickel(II), zinc(II), and cadmium(II) has been studied, and the highest affinity and selectivity was suggested to occur in the nickel(II) complexes [113]. A significant modification of the peptide conformation was also found in the nickel(II) complex of the dodecapeptide (HGGGHGHGGGHG) and the formation of both mono- and dinuclear species with nickel(II) coordinated to the imidazole and amide nitrogen Met. Ions Life Sci. 2, 63–108 (2007)


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atoms was suggested [114]. Moreover, the significance of the Ni(II)–Nim bonds is now evident from a wide range of methods used in molecular biology, which are based on the ability of nickel(II), when chelated to nitrilotriacetate, to selectively bind proteins containing stretches of consecutive histidine residues [115]. The complex formation processes of sulfhydryl-containing peptides have also been the subject of several reviews [16,17,21]. It is clear from these studies that nickel(II) has an outstanding affinity to bind the peptides of cysteine, but the stoichiometry and the binding modes of the species largely depend on the location of the thiol functions in the molecule. It has also been demonstrated that the thiol group can be an efficient anchor for the deprotonation and coordination of amide groups in nickel(II) complexes. Details of the versatility of the complex formation processes of simple cysteine containing peptides can be found in previous reviews [16,21]. In the last few years the investigations on the metal complexes of thiolcontaining peptides were mainly promoted by the biological significance of the zinc finger proteins. Nickel(II) is often used as a model for the spectroscopic characterization of zinc(II) in complexes. However, it should be noted that nickel(II) complexes of multiple thiolate donors reveal a great structural variety because the formation of both tetrahedral and square planar four-coordinated and octahedral six-coordinated species was suggested [116]. The formation of polynuclear species is another common feature of the complex-formation processes of nickel(II) with the peptides of cysteine. Accordingly, the presence of both mono- and polynuclear species has been reported in the nickel(II)-Ac-CysGlyCys-NH2 system and the dinuclear mixed ligand derivative was suggested as a promising model of the catalytic site of acetyl coenzyme A synthase [117]. Cyclopeptides containing cysteinyl residues provide a high structural variety for complexation, but they also make possible the formation of the NiII(S–Cys) 4 chromophore which can be considered as being the simplest structural motif of several metalloenzymes including [NiFe]-hydrogenases [118]. The peptides containing both thiolate and imidazole functions are especially efficient and interesting ligands for complexation with nickel(II). The formation of a square planar nickel(II)-thiolate bonded species was suggested with the N-terminally blocked form of -Cys-Xaa-His- peptides, while a pH and dioxygen-dependent coordination behavior was observed with the terminally free counterparts [119]. Glutathione (γ -GluCysGly) is one of the most common small biomolecules, present in cells of all living organisms at millimolar concentrations. Most reviews cited in Section 1 of this chapter give a brief account on the coordination chemistry of glutathione, while the most recent results were summarized in 1999 [120]. The great versatility of the complex-formation reactions of glutathione comes from the presence of the γ -peptidic bond between Glu and Cys residues, which separates the amino terminus from the amide and thiol functions. As a consequence, both the α-amino acid-like binding sites at the N-terminus and the thiol group in the side chain can be the anchoring site for metal ion coordination. In the case of nickel(II) both types of interactions are favored, resulting in the formation Met. Ions Life Sci. 2, 63–108 (2007)

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of several different species depending on pH and metal ion to ligand ratio. Most publications agree that coordination of glutathione to nickel(II) starts at the N-terminus via the formation of the five-membered (NH2, COO) chelate in the octahedral species. This binding mode is, however, followed by the formation of Ni(II)–S(thiol)-bonded complexes and it can induce either the deprotonation of neighbouring amide functions or the formation of sulfur-bridged polynuclear species in the form of diamagnetic square planar complexes [121–125]. The anchoring capability of the thiol function for amide deprotonation was also suggested for the nickel(II) complexes of α-mercaptopropionylglycine [126]. On the basis of the high affinity of glutathione towards the divalent transition elements it was suggested that glutathione can play an important role in the tolerance towards nickel toxicity [127]. On the other hand, more and more experimental findings support the view that nickel(II) accelerates the air oxidation of glutathione and that this has a significant impact on nickel carcinogenesis [128–131].

3.4.

Complexes of Peptide Fragments from Histones, Protamines, and Other Biologically Important Proteins

Metal ions can be bound by a large number of proteins. Such protein–metal interactions occur within specific amino acid residues called metal-binding motifs. The motifs that putatively bind metal ions like Ni(II) preferentially contain clusters of Cys and/or His [132]. Nickel compounds are well-established as human carcinogens [133], but the responsible molecular events remain to be fully understood. DNA binds Ni(II) only weakly [134], leaving nuclear proteins as possible targets for Ni(II). Particularly, the abundance of histones inside the cell nuclei makes them the primary target for metal ions. The nucleosome is composed of a histone octamer (containing two copies of H2A, H2B, H3 and H4 histones) and 145–147 base pairs of DNA bundled around it [135]. Available information on the binding modes of Ni(II) to proteins [136] and data for nickel–peptide complexes [17] indicate that the imidazole of histidine and thiol of cysteine could be thermodynamically preferred by Ni(II) among the donor groups provided by protein-building amino acids. H1 histone does not have any histidine or cysteine residue. However, inspection of the available histone sequences revealed several histidine and cysteine residues in H2, H3, and H4. Interactions of the TESHHK hexapeptide fragment representing the 34-amino acid residue C-terminus of histone H2A, resulted in hydrolysis of the peptide bond between the Glu and Ser residues under physiological conditions and formation of the albumin-like square planar Ni(II) complex with the SHHK sequence [137,138]. Studies on Ni(II) coordination with the N- and C-terminal protected hexapeptides TESHHK, TASHHK, TEAHHK, TESAHK, and TESHAK [139,140], in which Ala was inserted instead of Glu, Ser, or His-4 and His-5 in the TESHHK motif, were also performed. While substitution of His-4 or Glu by Met. Ions Life Sci. 2, 63–108 (2007)


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Ala did not affect the hydrolysis reaction, substitution of the His-5 or Ser residues inhibited the reaction, supporting the important role of the last two residues in peptide bond hydrolysis [139,140]. Above pH 7 SHHK and SAHK (hydrolysis products) coordinate to Ni(II) equatorially through the imidazole of His-3, the N-terminal amino group, and the two amide nitrogens located between Ser-1 and His-3, {NH2, 2N, Nim}, forming 4N albumin-like square planar complexes [141]. Spectroscopic evidence and theoretical predictions suggest that the positioning of the free imidazole ring, in the Ni-SHHK complex, above the coordination plane, induces the extra stability of the complex [142]. The coordinating properties of the N- and C-protected peptide fragment of the C-terminal domain (102–107), Ac-Glu-Leu-Ala-Lys-His-Ala-amide, of histone H2B towards Ni(II) ions were studied [143]. Imidazole was proposed as an anchoring site and at high pH the diamagnetic species with a {Nim, 3N} coordination mode was suggested. The peptide sequence (110–113), Cys-Ala-Ileu-His, of the H3 histone is evolutionarily strictly conserved among animal species and it contains the very attractive set of Cys and His residues as far as metal ion coordination is concerned [144]. This sequence was suggested to be a potential binding site for Zn(II), resembling to some extent the zinc finger binding pattern [145], but it seems that no study follows this suggestion. Cys-110 is the only free thiol in H3, and as such has often been employed as a chemical labeling site [146]. The peptide CAIH as a minimal model for the H3 histone was synthesized allowing metal-ion-binding studies and thus, its complex formation with Ni(II) was characterized [147]: At the physiological pH, Ni(II) and the peptide yielded unusual macrochelate complexes, in which Ni(II) was bound through Cys and His side chains in a square planar arrangement. Histidine (His-18) is also located in the histone H4 N-terminus that extends from the protein core, where it is accessible. Histone H4 is one of the most conserved proteins in nature, also for the N-terminal region (residues 1–22) [148]. This region features three repetitions of the sequence Gly-Lys-Gly and the unusual string of five basic residues, KRHRK. It was found that nickel is a potent inhibitor of histone H4 acetylation in yeast and in mammalian cells [149]. Studies with a minimal coordinating model of the H4 tail, AKRHRK [150], and of the larger N-terminal domain, SGRGKGGKGLGKGGAKRHRKVL (residues 1–22) [151,152], were performed. The spectroscopic data obtained for the Ni(II)– AKRHRK system have shown that histidine acts as an anchoring metal-binding site. The stability constant of the Ni(II) species with the H4 fragment was distinctly higher than that of the for Boc-AGGH peptide [150]. This may indicate that the positively charged side chains of Lys and Arg increase distinctly the stability of the 3N, 4N complexes [153]. Although the binding of Ni(II) in the physiological pH range is not very effective, the hydrophobic environment in the entire protein is expected to enhance the metal-binding capabilities, due to the multiple nonbonding interactions stabilizing the complex formed [154–156]. Met. Ions Life Sci. 2, 63–108 (2007)

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Protamines are small basic proteins which provide compact DNA binding in vertebrate sperm. Mammals possess two classes of protamines, P1 and P2. P1 rich in arginine and cysteine, but not histidine, is expressed in all mammals, while P2, which also contains histidines, has been detected in just a few mammalian species, including mice and humans [157,158]. Its presence at the levels of 50–70% of total protamine is, however, required for male fertility in humans [159]. HP2 contains the N-terminal tripeptide albumin-like sequence Arg-ThrHis. Analogous N-terminal sequences, containing histidine in the third position (Xx-Yy-His), are known for their specificity for Cu(II) and Ni(II) binding [18]. They are also present in several human peptide hormones and proteins, human serum albumin (HSA) being the most prominent [160]. The Asp-Ala-His sequence of HSA is the physiological carrier of Cu(II) [161], and it is also involved in nickel toxicity, providing the antigen for nickel allergy when coordinated to Ni(II) [162]. These facts strongly suggested that HP2 may contain a physiologically relevant Cu(II) and Ni(II) binding site at its N-terminus. The potentiometric and spectroscopic results [153] indicate that the N-terminal tripeptide motif Arg-Thr-His is the exclusive binding site for nickel ions at the metal to HP21–15 molar ratio not higher than 1. A solution structure of the Ni(II) complex with the N-terminal pentadecapeptide of human protamine HP2 (HP21–15) was elucidated with the use of one- and two-dimensional 1H NMR techniques and molecular modeling [163]. Cap43 is a novel gene induced by a rise in free intracellular Ca2 following nickel exposure [164,165]. No other metal compound significantly induced expression of this gene, indicating that it was expressed with a marked specificity to Ni(II) exposure [166]. Cap43 has a mono-histidine 10-amino acid-residue fragment (Thr-Arg-Ser-Arg-Ser-His-Thr-Ser-Glu-Gly) which is repeated three times in the C-terminal domain. It should be mentioned that such mono-histidine fragments, e.g., the octapeptide repeated regions in prion proteins, play a critical role in metal metabolism using a set of His imidazoles as the binding sites for metal ions [167]. The tetradecapeptide containing one 10-amino acid repeated sequence was analyzed for Ni(II) binding [168]. The 20-(Ac-TRSRSHTSEG-TRSRSHTSEG) and the 30-(Ac-TRSRSHTSEG-TRSRSHTSEG-TRSRSHTSEG) amino acid sequences were analyzed [169] as well. The 20-amino acid peptide can bind one or two metal ions, while the 30-amino acid fragment, binds up to three metal ions. Fibrinogen is a dimer, the two halves being held together by disulfide bridges, and fibrinopeptides A and B, together with fibrin, are produced by the cleavage of fibrinogen by thrombin when there is some injury to the body. Once the fibrinopeptides have been released the fibrin residues form a clot by side-to-side and end-toend aggregation [170]. Human fibrinopeptide A is a peptide containing 16 amino acid residues, Ala-Asp-Ser-Gly-Glu-Gly-Asp-Phe-Leu-Ala-Glu-Gly-Gly-Gly-ValArg. Local concentrations of fibrinopeptides near the site of injury will be high and they would be competitive with other biopeptides in interaction with metal ions, particularly Cu(II). Complexes with Ni(II) of the amino-terminal tetrapeptide fragment of human fibrinopeptide A (Ala-Asp-Ser-Gly) have been studied [86]. Met. Ions Life Sci. 2, 63–108 (2007)


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The major difference compared to that of tetraalanine is the greater importance of the NiH1L complex in the case of Ala-Asp-Ser-Gly. This suggests the formation of the Ni(II) chelation through the amino, amide, and β-carboxylate donor set. Arginine8-vasopressin (AVP) and arginine8-vasotocin (AVT) are naturally occurring neurohormones with similar structural features, each possessing a 20-membered ring linked by a disulfide bridge with a tripeptide tail. Spectroscopic studies have shown that the similarities in their primary structure are accompanied by similar peptide backbone conformations, as well as the conformations around the disulfide bridge [171,172]. The complexes of vasopressin and vasotocin with Cu(II) are one of the most stable Cu–peptide complexes with 4N coordination yet reported [95]. The major stabilization factor is the favorable positioning of the binding donor atoms within the ring formed by the disulfide bridge of the peptide. Secreted protein, acidic and rich in cysteine (SPARC) is a matricellular calciumbinding glycoprotein, which mediates the cell–matrix interactions without having any primarily structural role [173,174]. It has been reported as a product of some tumors and this result has been associated to the SPARC activity in angiogenesis [175]. The human protein consists of 286 residues divided into three distinct domains [176]. The second domain is a Cys-rich, follistatin-like (FS) domain (residues 53-137), in which all Cys residues are disulfide-bonded, with a N-linked carbohydrate moiety at Asn 99. This domain is characterized by the presence of two copper-binding sites, the strongest of which contains the sequence LysGly-His-Lys (KGHK) (residues 120–123). Both KGHK and the longer peptides containing this sequence have been recognized to regulate angiogenesis in vitro and in vivo [177]. The tetrapeptide KGHK binds Ni(II) ions in a tetradentate albumin-like fashion {NH2, 2N, Niim}. Angiotensin II, a peptide hormone which has the sequence Asp-Arg-Val-TyrIle-His-Pro-Phe, is involved in the regulation of blood pressure and has been shown to interact with metal ions in biological systems [178]. The study of the complexes of angiotensin II and two of its peptide fragments, Asp-Arg-Val-Tyrand MeCO-Tyr-Ile-His, with Ni(II) shows that metal ions at high pH form 4N species with the metal ion bound at the N-terminus of angiotensin II, giving a complex closely similar to that formed by Asp-Arg-Val-Tyr [85]. There may be a role for the imidazole site at lower pH range, but comparison of the Ni(II) complexes of MeCO-Tyr-Ile-His with those of Asp-Arg-Val-Tyr shows the latter peptide to be more effective in 4N complex formation. The results suggest that, with Ni(II), the terminal amino nitrogen of the Asp residue is a more effective center to initiate coordination than is the imidazole nitrogen. The thyrotropin releasing factor (TRF, L-pyroglutamyl-L-histydyl-L-prolinamide, Pyr-His-Pro-NH2) and melanostatin (MIF, L-prolyl-L-leucyl-glycinamide, Pro-Leu-Gly-NH2) are both oligopeptide hormones containing among others a proline residue at the C- and N-terminal ends, respectively. The interactions of Ni(II) with Pyr-His-Pro-NH2 and the Pyr-His dipeptide analog have been studied [179]. In both systems, Ni(II)-TRF and Ni(II)-Pyr-His, at least two square planar Met. Ions Life Sci. 2, 63–108 (2007)

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complexes are formed with the metal-coordination sites at N3 imidazole, N of the peptide linkage, and the deprotonated nitrogen of Pyr. The difference between these complexes is the pyrrole type nitrogen (N1) of imidazole, which may be protonated (pH 10) or deprotonated (pH 10). The drastic variations in the CD spectra in the d–d transition region due to deprotonation of the N1 imidazole, result most probably from a change in the chelate ring conformation. The N1 deprotonation of the imidazole ring changes its aromatic character and the coordinated N3 site alters its π acceptor properties. This may change the Ni(II)–N3 bond length slightly which is very sensitive to the geometry of the metal-imidazole interaction [180]. Melanostatin is a hypothalamic hormone and a therapeutic agent for Parkinson’s disease [181]. Complex formation between Ni(II) and MIF was found to be exceedingly slow at 25C, and 1:1 metal:ligand solutions tended to precipitate [58]. Above pH 8 a NiH3L complex could be identified clearly as the planar complex. It is therefore reasonable to assume that this is a 4N complex comparable to the Cu(II) analog [182,183]. The gonadotropin-releasing hormone (GnRH) or luteinizing-hormonereleasing hormone (LHRH) is a decapeptide (pEHWSYGLRPG) and plays an essential role in mediating the neuroendocrine control of reproductive processes. It was shown [184] that Cu(II), Ni(II) and Zn(II), may distinctly change the biological activity of the LHRH, as was found in the case of TRF [185]. The complex of Cu(II) with GnRH brought about a high release of LH and an even higher release of FSH. The Ni(II) complex showed a similar although less distinct effect [186]. In the study of metal–GnRH complexes interacting with the rat pituitary receptor [187], the Cu(II) complex was shown to act more efficiently than native GnRH, whereas the activity of nickel or zinc complexes was slightly lower. The coordination of Ni(II) starts at a pyridine-like nitrogen and then adjacent amide nitrogens complete the coordination around the metal ion forming 3N of {Nim, 2N} and 4N {Nim, 3N} coordinated species [188,189]. The studies involving several metal ions have shown that LHRH is very specific ligand for Ni(II) and the coordination mode found for this metal ion is considerably different from that occurring for Cu(II). 1H NMR measurements have been performed for the free GnRH and its complexes with Ni(II) in DMSO (dimethyl sulfoxide) [190]. The examples mentioned above clearly indicate that Ni(II) ions are critical factors for oligopeptide structures and their biological activities. It is actually very likely that many natural oligopeptides are affected by metal ion coordination in their biological functions.

3.5. Redox and/or Catalytic Reactions Involving Nickel(II) Complexes Peptide complexes of Ni(III) exhibit a variable stability in solution, persisting from seconds to hours [191–194]. Nickel(III) complexes undergo self-oxidation–reduction, Met. Ions Life Sci. 2, 63–108 (2007)


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O (–) O H2N

O

O

(–)

H

OH2

(–) N

O k1obsd

NiIII H2N

N O (–)

O N

H+ (–)

O

NH2 NiIII

N H2

NiII(aq) + peptide fragments

N O OH2

O

k2obsd

H O (–) O

(A)

(B)

(C)

Scheme 4. Reprinted from [80] with permission from the American Chemical Society.

catalyzed by either acid or base [192,195] in which the ligand is oxidized and the metal ion is reduced [196]. Oxidation of blue [bis(dipeptide)Ni(II)] yields violet–black solutions of Ni(III) complexes which are long-lived species in neutral solution. The acid-catalyzed conversion of the violet–black complex (A) to the yellow species (B) is followed by the acid-independent redox reaction to give diamagnetic products (C) (Scheme 4; reproduced by permission from [80]) [6]. The studies of the redox decomposition of Cu(III) and Ni(III) tripeptide complexes have shown that amino acid residues with methyl groups on the α-carbon tend to stabilize the trivalent oxidation state. Furthermore, the residue in the carboxyl terminal is oxidized preferentially in the order Gly Ala Aib. Oxidation of the orange low-spin [NiII(H1Aib2)2] 2 complex gives the dark olive-green tetragonally compressed [NiIII(H1Aib2)2] that is very stable in neutral and basic solutions [80]. In perchloric or dichloroacetic acid solution (pH 0.3–2.4), the tetragonally compressed [NiIII(H1Aib2)2] complex (A) decomposes to give Ni(II) plus peptide and oxidation products (C) in a three-reaction sequence (Scheme 5; reproduced by permission from [80]). Acid reacts with [NiIII(H1 Aib2)2] according to A → B → B → C to give two other Ni(III) complexes before decomposition to Ni(II) occurs. Addition of tripeptide excess (L) to solutions of [(tripeptido)Ni(III)], [NiIII(H2L)], gives [bis(tripeptido)Ni(III)] complexes. The five-nitrogen complex [Ni(H2L)(H1L)] 2 converts to a six-nitrogen tetragonally elongated species, [NiIII(H2L)2]3, above pH 11 [197]. When solutions of [bis(triglycinato) Ni(III)] complexes are made basic, a decoloration occurs in which the Ni(III) Met. Ions Life Sci. 2, 63–108 (2007)

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KOZLOWSKI et al. kAB

O

H2N

O

(–)

(–)

N

O

H2N

(–)

O

H2N

N

O

O

(A)

O

(B')

H2 N

N NiIII

k B'B

O

O

O OH2

O

O (–)

(–)

H

(–)

NiIII

kAB'

N

(–)

O

O

N

O NiIII

OH2

N H2

kBC

N OH2

(+) NH3

O H O

NiII(aq) + peptide

(–)

O

(B)

(C)

Scheme 5. Reprinted from [80] with permission from the American Chemical Society.

slowly oxidizes the peptide. The main site of peptide oxidation is the carboxylate terminal peptide residue. The reduction of Ni(III) peptides is accompanied by a release of coordinated water molecules: [ Ni III (Hn L)(H 2 O)2 ] e [ Ni II (Hn L)] 2H 2 O L is the peptide ligand and n is the number of deprotonated amide nitrogens in the complex. The Ni(III)/Ni(II) reduction potentials vary from 0.79 to almost 0.9 V versus the normal hydrogen electrode, depending on the nature of the peptide [198]; hence, in aqueous solution, Ni(III)–peptide complexes are moderately strong oxidants. The values of the entropy difference between oxidation states, SoII–SoIII are positive for the nickel couples and correspond to a loss of two water molecules for the reduction of Ni(III) peptide complexes [199]. The d8 electron configuration of Ni(II) behaves like that without axial coordination of water, while the d7 electron configuration of Ni(III) behaves like the one with two water molecules coordinated axially. Little is known about the rates of electron transfer in the Ni(III)/Ni(II)– peptide complexes. The kinetics of reduction by iodide involve axial coordination of iodide followed by the reduction of Ni(III) to Ni(II) [200]. Deprotonated peptide complexes of nickel(II) and nickel(III) have electron self-exchange rate constants that depend considerably on the structure of the complex [201]. The electron transfer reactions are catalyzed by bridging ligands such as Br and Cl, which can coordinate Ni(III) but are inhibited by non-bridging ligands like pyridine [201]. An electron transfer self-exchange rate constant was measured for the [Ni III (H2Aib3)(H 2O) 2] and [Ni II (H2Aib3)] couple, where Aib3 is the tripeptide of α-aminoisobutyric acid [202]. The rate of electron transfer between Ni(III) and Ni(II) decreases when pyridine is present in Met. Ions Life Sci. 2, 63–108 (2007)


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the solution. The axially coordinated pyridine blocks the one water-bridging pathway, but still permits the Ni(III)–H2O–Ni(II) bridge from the opposite side of the complex. Normally, Ni(II) is not easily oxidized, but is activated by peptide coordination so that it reacts with molecular oxygen and catalyzes the oxidation of the ligand. Tetra- and penta-peptide complexes of Ni(II) consume O2 in neutral solutions as the metal ion catalyzes the oxidation of the peptide to give a number of products [203]. Potentiometric studies have shown that [NiII(H2GGH)] is the major species present in millimolar aqueous solutions of NiII and GGH at pH values above 6.4 [204]. This complex is known to be highly sensitive to O2 in aqueous solution [191,205]. By electrochemical studies, electronic absorption and electron paramagnetic resonance (EPR) spectroscopy it was confirmed the reaction involves a Ni(III) intermediate, and it was established by X-ray crystallography that the Ni(II)–peptide product is decarboxylated and hydroxylated at His Cα [206]. Formation of Ni(III) is a common feature in oxidations of square planar Ni(II)–peptide complexes [192]. The crystal structure of [Ni(II)GlyGlyα-OH-Hm] (where Hm is histamine) shows a square planar complex, chelated by the amine terminal nitrogen, two deprotonated peptide nitrogens, and a histidyl imidazole nitrogen. The decomposition products of [NiIII(H2Gly2HisGly)] in neutral and basic solution [207] as well as the decomposition kinetics of the Gly2HisGly and Gly2Hm complexes of Ni(III) [208] were recently studied. Self-decomposition of the Ni(III)–peptide complex of Gly2HisGly occurs by base-assisted oxidation of the peptide. At pH 7, the major pathway is a four-electron oxidation (via 4 Ni(III) complexes) at the α-carbon of the N-terminal glycyl residue. In another major pathway above pH 7, two Ni(III)–peptide complexes coordinate via an oxo bridge in the axial positions to form a reactive dimeric species. This dimer generates two Ni(II)–peptide radical intermediates that cross-link at the α-carbons of the N-terminal glycyl residues. These species are formed via an oxo-bridged Ni(III)–peptide dimer which is very reactive. A [(NiIIIsalen)2O] species has been isolated and its crystal structure determined [209]. The self-decomposition reaction of [NiIII(H2Gly2Hm)] at pH 5 to 7.0 is a two-electron oxidation at the N-terminal glycyl residue to give [NiII(H2glyoxylglycylhistamine)] [210]. By contrast, the [NiIII(H2Gly2HisGly)] complex undergoes a four-electron oxidation at the same site. Above pH 8.5, [NiIII(H2Gly2Hm)] more closely mimics the reactivity of [NiIII(H2Gly2HisGly)] by forming an oxo-bridged Ni(III) dimer intermediate that reacts to form a cross-linked peptide product. The peptide– peptide cross-linking Ni(III) species is not found in the Cu(III) analogs [211], as the square planar geometry of Cu(III) precludes the formation of an oxo bridge between two Cu(III) species. The ‘uncomplexed’ Ni(II) does not activate H2O2 [212,213] and it is only weakly bound and generally innocent for double-stranded DNA [214,215]. On the other hand, the effects of Ni(II) on DNA in vivo suggest Fenton-like behavior Met. Ions Life Sci. 2, 63–108 (2007)

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[216]. In chemical studies on potential ligands that might facilitate nickel’s oxidative activity it has been established that only low-spin Ni(II) complexes (mainly of square planar structure) can be oxidized to Ni(III) in aqueous solution [217]. Quite interestingly, this is a common feature for the Ni(II) complexes with oligopeptides, even as simple as tetraglycine [192]. However, for most of the peptides, the square planar complexes are formed at high, nonphysiological pH values above 9 [17]. The relevant exclusion from this rule is provided by peptides having a His residue at position 3, which form low-spin, square planar complexes at physiological pH [18]. The simplest of these peptides, Gly-Gly-His, forms a complex with Ni(II) that is able to activate not only hydrogen peroxide [218], but also molecular oxygen in ambient air [191]. Ni(II) complexes with Gly-Gly-His and its analogues Xaa-Yaa-His [219,220] and DNA binding proteins, extended N-terminally with the Gly-Gly-His sequence have gained interest as potential specific DNA [221–223] or protein [224,225] cleavage agents in the presence of oxidants. Ni(II)-Xaa-Yaa-His metallopeptides are unique in their ability to incorporate and position within a metal complex framework the same chemical functionalities used by proteins and natural products for the molecular recognition of DNA and RNA [226,227]. The metallopeptides activated by KHSO5 (oxone), MMPP (magnesium monoperoxyphthalate) or H2O2 may be involved in modifications of DNA, RNA and proteins. The reactive species is proposed to be a high-valent, peptide-bound NiIII–HOᠨ or NiIVO which is generated from the heterolytic splitting of the oxygen–oxygen bond present in KHSO5, MMPP, and H2O2 [228,229]. Further studies revealed that the natural histidyl oligopeptides carnosine, homocarnosine, and anserine are capable of enhancing oxidation of free 2-deoxyguanosine (dG) to 8-hydroxy-2-deoxyguanosine (8-OH-dG) by H2O2 in the presence of Ni(II) [230]. A similar reaction was observed for the Ni(II)-tetraGly system [231]. Histidine-containing Ni(II) oligopeptides catalyze the disproportionation of hydrogen peroxide in a process which generates active intermediates capable of hydroxylating p-nitrophenol and oxidizing uric acid to allantoin. An oxene moiety, namely NiO2 is postulated as the active species in these H2O2dependent reactions [232]. L-Histidine, L-cysteine, reduced glutathione (GSH) and other bioligands, which are ubiquitously present in biological systems, are recognized as antioxidants. Studies have shown that Ni(II) complexed with these ligands catalyzes the disproportionation of H2O2, leading to the generation of hydroxyl radicals [131]. Amino acid binding domains for Ni(II) were identified in the core histones H3, H2A, and H4 and in protamine P2. Some of the resulting metal complexes did, in fact, promote DNA base oxidation [137,138,153,233,234]. In one of these complexes, Ni(II)-H2A8 (A8 SHHKAKGK, fragment of histone H2), nickel was capable of significantly enhancing oxidation of the peptidic ligand.

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3.6. Nickel(III) Peptide Complexes Nickel(II) promotes deprotonation of peptide and amide nitrogens upon its coordination to oligopeptides [16–18]. Coordination of deprotonated peptide nitrogens stabilizes the trivalent oxidation state of nickel. The Ni(III)–peptide complexes are readily obtained from the corresponding Ni(II) complexes by chemical or electrochemical oxidation [191,198]. These Ni(III) species are moderately stable in aqueous solution. Standard electrode potentials from 0.96 to 0.79 V were determined for a series of Ni(III)/Ni(II)-peptide couples in aqueous solution. The values of Eo decrease with the number of bulky C-substituents, depending upon their position in the amino acid residues. The increase in Eo for Gly-Gly-Val compared with GlyGly-Gly can be attributed to the interaction of the bulky substituent with axial solvation as it was proposed in the case of the nickel macrocyclic complexes [235]. Ligands with histidine as the third amino acid residue have an amine nitrogen, two deprotonated peptide nitrogens, and one imidazole nitrogen bound to nickel. Although the imidazole group is higher in the spectrochemical series than the carboxylate one [236], the values of Eo of the histidine-containing peptides were larger than those of the tripeptide complexes. High potentials were also found for the same complexes of copper and were attributed to a combination of the relative effects of cumulative ring strain and π back-bonding [237]. UV-visible absorption and circular dichroism measurements for nickel tetraand pentapeptide complexes were performed [238]. For the [CuIII(H3L)] and [NiIII(H3L)] complexes one spectral feature is a charge-transfer band at 365 nm and 320 nm, respectively, with molar absorptivities of 4500–7500 M1 cm1, depending on the nature of the peptide [237]. Electron paramagnetic resonance was used to show that oxidation of Ni(II)– oligopeptide complexes results in removal of the electron from the metal center so that the unpaired electron is associated with nickel(III) rather than with a coordinated nitrogen or carbon radical. Oxidation of Ni(II)–oligopeptide complexes in aqueous solution yields paramagnetic products and EPR spectra are consistent with Ni(III) in the tetragonal geometry with four donor groups of the oligopeptide in the equatorial plane and two axially bound water molecules [193]. The unpaired electron is located in an orbital which has a large amount of dz2 character, although broadening effects attributed to hyperfine splitting from equatorial nitrogens are observed. The magnitude of the equatorial g value increases as the strength of the equatorial donor increases in the order N (peptide) NH2 imidazole COO [236]. In the case of Ni(III)-glycylglycylglycinamide, [NiIII(H3G3-NH2)], the nature of the tetragonal distortion was investigated using ammonia as a probe. As the concentration of ammonia in solutions of [NiIII(H3G3-NH2)] increased, two new paramagnetic species were detected. The EPR spectrum suggests a species which has a single nitrogen nucleus (I 1) bound in an axial position. It is rather unlikely for a nitrogen donor that Met. Ions Life Sci. 2, 63–108 (2007)

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is an equatorial peptide ligand to move to an axial position [239]. A second molecule of ammonia may be added to give the complex whose EPR spectrum indicates that two ammonia molecules are bound in the equivalent axial positions. The difference in the stability constants for species obtained after addition of the fifth and sixth nitrogen donors to [NiIII(H3G3-NH2)] reveals a strong preference for binding of five nitrogen donors. Indeed, the d7 Ni(III)-complexes show strong preference for the square pyramidal arrangement of donors. The square pyramidal geometry is also common with low-spin d7 Co(II) [240,241]. The EPR spectra of various histidine-containing tripeptides and of bleomycin with Ni(III) are consistent with a tetragonal geometry (gxx, gyy gzz) rather than a square planar geometry (gzz gxx, gyy). The Ni(III) complexes with sulfhydryl groups in equatorial positions exhibit larger gxx values and more rhombic symmetry than those with the amino ligands. The large gxx value of the sulfur–nickel(III) complexes suggests the trend of S NH2 in the equatorial donor strength for Ni(III) [242]. By using an electrochemical method stability constants for axial coordination of monodentate nitrogen ligands to nickel(III)–tripeptide complexes in aqueous solution were determined [243]. The stability constants at 25C decrease in the order imidazole NH3 N 3 pyridine and are relatively insensitive of the nature of the peptide. The axial coordination to Ni(III) is unhindered, even when the α-carbons in the peptide chain are fully substituted. Substitution in the second axial site is not favored and was not detected at room temperature, but it occurs in frozen aqueous solutions with ammonia. The triglycine, Gly-Gly-Ala and Gly-Ala-Gly, complexes of Ni(III) react with bpy, phen, en, and dien to form ternary complexes where the added ligands are chelated as it was seen from EPR spectra [244]. The complexes [NiIII(H2GlyGly-Ala)(H2O)2] readily forms mixed-ligand complexes with en and dien. These polyamine ligands are able to displace the axially coordinated water molecules as well as the carboxylate oxygen of the tripeptide complex. The frozen-solution EPR spectra are indicative of tetragonally elongated structures, with gxx gyy gzz. Peptide complexes with Aib inserted as the third residue are sterically hindered in the formation of chelate adducts. The steric restrictions that arise from the two methyl groups on the third peptide residue of the Aib3 ligand limit access to the equatorial site where the carboxylate oxygen is coordinated. Hence, en and dien are unable to displace the carboxylate oxygen and form a chelate ring. EPR spectra of frozen aqueous solutions show that the [NiIII(H2GGA)(dien)] complex exists in three forms that occur as a function of increasing pH. This indicates a tridentate coordination of dien. Formation of Ni(III)–tripeptide axial donor species leads to complexes that are more stable than the parent Ni(III) complex with regard to redox decomposition in neutral and basic solutions [244]. Nickel(III) complexes of tripeptides and dipeptide–Schiff bases studied by cyclic voltammetry indicate that, depending upon the nature of the coordinated groups and the structure of fused ring systems, the electrode potentials of Ni(III)/ Met. Ions Life Sci. 2, 63–108 (2007)


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Ni(II) couples vary in the range 0.7–1.4 V [245]. The stronger donating groups such as amino and deprotonated amide nitrogens assist the stabilization of metal ions in higher oxidation states. The trivalent nickel with a smaller ionic radii is stabilized by the 5-5-5, but not by the 5-6-5 or 6-5-6 fused ring structure as is the case in bivalent nickel complexes. Ni(II) forms blue complexes with two dipeptide molecules, and for the case of glycylglycine the geometry has been shown to be a tetragonally compressed octahedron [74,239]. Electrochemical or chemical oxidation of the Ni(II) species results in the formation of a violet–black paramagnetic complex with the proposed formula [NiIII(H1GlyGly)2]. The frozen EPR spectrum for [NiIII(H1GlyGly)2] corresponds to that of a nonaxially symmetric complex with one unpaired electron. The spectrum has g g⊥, in contrast to the typical situation for Ni(III) peptide complexes, where g⊥ g for a tetragonally elongated octahedral distortion. Similar EPR spectra were obtained for all the [bis(dipeptide)Ni(III)] complexes (dipeptides with glycyl, alanyl, and α-aminoisobutyryl residues). In the case of the bis(dipeptide) complexes the amine nitrogens, the deprotonated peptide nitrogens, and carboxylate donors prefer a compressed octahedral environment with nickel(II) [74] and cobalt(III) [246]. Addition of ligand excess (L) to solutions of [(tripeptido)Ni(III)], [NiIII(H2L)], gives [bis(tripeptido)nickelate(III)] complexes. In the initial reaction [NiIII(H2L)(L)] forms, but it is a transitory species that rapidly loses a proton to give [NiIII(H2L)(H1L)] 2, a complex with five nitrogens coordinated to nickel [197]. This five-nitrogen complex is relatively stable from pH 6 to 11. EPR studies frequently are used to propose the structure of Ni(III) species because Ni(III)-peptide complexes are too unstable to obtain crystals for X-ray crystal structure determinations [6,193,235,247,248]. For [NiIII(H2G3)(H1G3)] 2 three peaks in the g region indicate one axial nitrogen coordinated to the metal ion. At pH 13, five peaks in the g region [197] indicate that the second nitrogen is axially coordinated to nickel. The proposed formula is [NiIII(H2G3)2]3 with six nitrogens, four of which are deprotonated N-peptide groups, coordinated to Ni(III). Peptides that contain Aib in the third residue, GGAib, AAibAib and AibAibAib, do not form bis-complexes. Substitution of methyl groups for methylene hydrogens tends to increase the stability of peptide complexes; e.g., Ni(III) and Cu(III) complexes of GlyGlyAla are significantly more stable than their triglycine analogs [249]. Oxidation of the [Ni(II)(Aib2)2] complex gives an olive-green [NiIII(H1Aib2)] complex, which is indefinitely stable in neutral solution and decomposes only slowly, even in strong base [80]. The relative rates of the self-redox decomposition of the [bis(dipeptido)Ni(III)] complexes decrease in the series GG AG AibG GA AA Aib2. The EPR spectrum for the [NiIII(H1Aib2)2] complex shows g g⊥, in contrast to typical EPR spectra of Ni(III)–tripeptide complexes with a tetragonally elongated octahedral geometry with g⊥ g. This spectrum is similar to that obtained for [NiIII(H1G2)2] [6]. Met. Ions Life Sci. 2, 63–108 (2007)

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Ni(III)–peptide complexes with histidine and histamine as the third residue are relatively stable in acidic solution with half-lives of more than 10 h [208]. However, in basic solution these Ni(III)–peptide complexes are unstable, and the loss of Ni(III) is complete within milliseconds to seconds. The deprotonation of the axially coordinated water molecule was not observed below pH 11 with non-histamine/histidine-containing Ni(III) complexes [243,197]. Murray and Margerum [243] estimated the pKa value of the coordinated water molecule to be greater than 11 based on reduction potential measurements of the Ni(III)– peptide complexes in the pH range 6.5–10.5. Similarly, Kirvan and Margerum [197] found that the pKa value of a coordinated water in [NiIIIAla3] is about 11.3.

4. FORMATION OF NICKEL(II) COMPLEXES UNDER BIOLOGICAL CONDITIONS: MODEL CALCULATIONS IN MULTICOMPONENT SYSTEMS The relative stabilities of the various chelates built up from N- and O-donor ligands have already been compared in Figure 1. It was clear from these plots that the five-membered (NH2, NH2) and six-membered (NH2, Nim) chelates predominate over the common (NH2, COO) coordination modes of amino acids in the neutral and slightly basic pH ranges. The differences in the complex-forming abilities of various amino acids are even more evident in Figure 2, where the metal ion 1.0

His complexes Ni 2+

Fraction of Ni(II)

0.8

Cys complexes

[Ni(Cys)2]2– [Ni(His)] + [Ni(His)2]

0.6

0.4

0.2

Gly complexes [Ni(HCys)]+

0.0 3

4

5

6

7

8

9

10

pH 11

Figure 2. Concentration distribution of the complexes formed in the nickel(II)–glycine–histidine–cysteine 1:3:3:3 (mmol) system. Stability constants are taken from [22] (glycine), [31] (histidine) and [250] (cysteine). Met. Ions Life Sci. 2, 63–108 (2007)

Ni COMPLEXES OF AMINO ACIDS AND PEPTIDES 1.0

95

complexed Cu(II) [CuA 2H 2]2+

Fraction of His

0.8

0.6 [CuA 2] complexed Ni(II)

0.4 [CuA]

H 2A+

0.2

+

[NiA]+ [NiA2]

[CuAH] 2+ [CuA 2H]

+

[ZnA] +

complexed Zn(II)

0.0 3

4

5

6

7

8

9

10

pH 11

Figure 3. Concentration distribution of the complexes formed in the copper(II)– nickel(II)–zinc(II)–histidine 1:1:1:3 (mmol) system. Stability constants are taken from [31].

speciation of the Ni(II)–glycine–histidine–cysteine 1:3:3:3 system is plotted as a function of pH. It can be seen from Figure 2 that the amino acids histidine and cysteine bind Ni(II) almost exclusively at any pH. As a consequence, glycine and the other common bidentate amino acids cannot compete with histidine and/or cysteine for nickel(II) binding under biological conditions. It is also clear that histidine and cysteine have a comparable nickel binding affinity in the physiological pH range, in agreement with previous findings that these amino acids are the major metal-ion-binding sites in proteins. It is also important to compare the affinity of histidine and cysteine towards different transition elements. Because of the redox reaction between copper(II) and thiol ligands such a comparison cannot be easily demonstrated with cysteine, therefore Figure 3 reveals the copper(II), nickel(II), and zinc(II) binding affinity of histidine only. In agreement with the Irving–Williams order it follows the trend: Cu(II) Ni(II) Zn(II), but the pH dependence of the complexation with the three metal ions is also very significant. If one takes into account the different abundance of the three metal ions in biological fluids, the preference for copper(II) binding is even more pronounced, but it is also evident that all three metal ions can coordinate to histidine in the physiological pH range. The model calculations of multicomponent systems containing peptide ligands led to similar conclusions. It is not demonstrated by a speciation curve here, but GlyGly, similarly to glycine, cannot compete with the peptides of histidine and/or cysteine for nickel(II) binding. The peptide binding strength of zinc(II) Met. Ions Life Sci. 2, 63–108 (2007)

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1.0 Ni2+

[Ni(H –2AlaAlaCys)] 2–

0.8

Fraction of Ni(II)

AlaAlaCys complexes [Ni(H–2GlyGlyHis)]

–

0.6

0.4

GlyGlyHis complexes

0.2 [Ni(HAlaAlaCys)] +

0.0 3

4

5

6

7

8

9

10

pH

11

Figure 4. Concentration distribution of the complexes formed in the nickel(II)– GlyGlyHis–AlaAlaCys 1:1:1 (mmol) system. Stability constants are taken from [251] (GlyGlyHis) and [252] (cysteine).

is also much reduced compared with those of copper(II) and nickel(II) because zinc(II) ions are generally not able to promote deprotonation and coordination of amide functions. The tripeptides GlyGlyHis and AlaAlaCys are well known to form stable complexes both with copper(I) and nickel(II). Figure 4 demonstrates the differences in the nickel-binding capacity of GlyGlyHis and AlaAlaCys. The conclusions are rather similar to those obtained for the corresponding amino acids from Figure 2. Namely, the nickel(II) binding strength of the (NH2, N, N, Nim) and (NH2, N, N, S) coordination modes is comparable around pH 7, but the thiol-containing species are favored in alkaline solution.

5.

CONCLUSIONS

In summarizing the metal binding affinity of different amino acid residues it can be stated that the complexation with copper(II) is always favored among the 3d transition elements. The amino acids and/or peptides, however, are always present in a high excess compared with the metal ions under biological conditions, and as a consequence, nickel(II) can bind either to amino acids and peptides or proteins. Figures 2–4 clearly indicate that histidine-Nim and cysteine-S donor atoms are the major binding sites. Deprotonation and nickel(II) coordination of the peptide amide bond generally occurs only above the physiological pH range, but histidyl Met. Ions Life Sci. 2, 63–108 (2007)


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and cysteinyl residues in the appropriate locations can work as anchors for the promotion of this process. Up to now, there has not been an unambiguous proof for the existence of Ni(II)–Namide bonds in biological systems, but further studies on the nickel(II) complexes of peptides or derivatives of histidine and cysteine are justified by the high stability of Ni(II)–Nim and Ni(II)–Sthiolate bonds. Another important feature of the coordination chemistry of Ni(II)–amino acid/peptide complexes comes from the increasing number of publications on the catalytic role of nickel complexes and the easy formation of nickel(III)–peptide systems. Although the biological relevance of Ni(III) is implicated, e.g., for nickel superoxide dismutase (see Chapter 10) and nickel iron hydrogenases (see Chapter 7), its properties need to be further evaluated, yet the redox behavior of the Ni(II)/Ni(III) pair has become clear.

ABBREVIATIONS Aib AVP AVT BIMA Boc bpy CD Daba Dapa dG dien DMSO DOPA en FS FSH GGH GnRH GSH Hm HmS HSA im LH LHRH MIF MMPP

aminoisobutyric acid arginine8-vasopressin arginine8-vasotocin bis(imidazol-2-yl)methylamine t-butoxycarbonyl 2,2-bipyridine circular dichroism 2,4-diaminobutyric acid 2,3-diaminoproponic acid 2-deoxyguanosine diethylenetriamine dimethyl sulfoxide 3,4-dihydroxyphenylalanine 1,2-diaminoethane follistatin follicle-stimulating hormone glycyl-glycyl-histidine gonadotropin-releasing hormone reduced glutathione histamine α-hydroxymethylserine human serum albumin imidazole luteinizing hormone luteinizing-hormone-releasing hormone melanostatin magnesium monoperoxyphthalate Met. Ions Life Sci. 2, 63–108 (2007)

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phen pn PNA SPARC TRF

1,10-phenanthroline 1,3-diaminopropane peptide nucleic acids secreted protein, acidic and rich in cysteine thyrotropin-releasing factor

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23.

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Met. Ions Life Sci. 2, 109–180 (2007)

4 Complex Formation of Nickel(II) and Related Metal Ions with Sugar Residues, Nucleobases, Phosphates, Nucleotides, and Nucleic Acids Roland K. O. Sigel1 and Helmut Sigel 2 1

Institute of Inorganic Chemistry, University of Zürich, Winterthurerstrasse 190, CH-8057 Zürich, Switzerland

2

Department of Chemistry, Inorganic Chemistry, University of Basel, Spitalstrasse 51, CH-4056 Basel, Switzerland

1. INTRODUCTION 110 2. NICKEL(II)–SUGAR INTERACTIONS 112 2.1. Carboxyhydrate Amines: A Strong Primary Binding Site May Help in Hydroxyl Group Coordination 112 2.2. Ribose Hydroxyl Groups Are Poor Binding Sites in Complexes of Nucleoside Monophosphates 113 2.3. A Favorable Steric Setting and a Reduced Solvent Polarity May Promote Metal Ion–Hydroxyl or Carbonyl Group Binding 115 3. INTERACTIONS OF NICKEL(II) WITH NUCLEOBASE RESIDUES 118 3.1. Nickel(II) Complexes of Purine Derivatives 118 3.2. Nickel(II) Complexes of Pyrimidine Derivatives 122 4. COMPLEXES OF NICKEL(II) WITH PHOSPHATES 128 5. NICKEL(II) COMPLEXES OF NUCLEOTIDES 131 5.1. Some General Considerations 131 5.2. Complexes of Nucleoside 5-Monophosphates 132 5.2.1. Definition of the Equilibrium Constants 132 5.2.2. Properties of Pyrimidine-Nucleoside 5-Monophosphate Complexes 134 Metal Ions in Life Sciences, Volume 2 © 2007 John Wiley & Sons, Ltd


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5.2.3.

6.

7.

8.

9. 10.

1.

Properties of Purine-Nucleoside 5-Monophosphate Complexes 5.3. Complexes of Nucleoside 5-Di- and Triphosphates COMPLEXES OF SOME LESS COMMON NUCLEOTIDES 6.1. Complexes of 2- and 3-Nucleoside Monophosphates 6.2. Complexes of Orotidinate 5-Monophosphate 6.3. Complexes of Xanthosinate 5-Monophosphate 6.4. Flavin Mononucleotide Complexes COMPLEXES OF SOME NUCLEOTIDE DERIVATIVES AND ANALOGS 7.1. Complexing Properties of 1, N 6 -Ethenoadenosine and of Its Phosphates 7.2. Complexes of (N1)-Oxides of Adenosine and Inosine Nucleotides 7.3. Complexes of Nucleoside 5-O-Thiomonophosphates 7.4. Complexes of Acyclic Nucleotide Analogs 7.5. Nickel(II) Binding to Nucleotides Containing a Platinum(II)-Coordinated Nucleobase Residue MIXED LIGAND COMPLEXES CONTAINING A NUCLEOTIDE 8.1. Some General Comments and Definitions 8.2. Ternary Nickel(II) Complexes Containing ATP 4 and a Buffer Molecule 8.3. Mixed Ligand Complexes Containing a Nucleotide and a Further Monodentate or Bidentate Ligand: Release of N7 and Formation of Stacks NICKEL(II) BINDING IN NUCLEIC ACIDS CONCLUDING REMARKS ACKNOWLEDGMENTS ABBREVIATIONS AND DEFINITIONS REFERENCES

135 139 144 144 146 146 148 149 150 150 154 156 158 159 159 161

164 165 168 169 169 172

INTRODUCTION

The abundance of nickel in the Earth’s crust and in seawater is comparable to that of many other transition metal ions (see also Chapters 1 and 2 of this volume) with Met. Ions Life Sci. 2, 109–180 (2007)

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a long established history of biological importance [1]. However, when Mertz in 1970 reviewed the nutritional aspects of trace element research, he found that nickel is unique among the elements of the first transition series because all of its neighbors, i.e., V, Cr, Mn, Fe, Co, Cu, and Zn, had been reported to be essential or at least as having a particular biological function [2] but not nickel. Indeed, there have been explanations for the apparently virtual absence of nickel from living systems [3]. This situation changed dramatically in 1975 with the discovery of Blakeley and Zerner and their coworkers [4] that jack bean urease is a metalloenzyme containing nickel. Indeed, research on the role of nickel in biology exploded dramatically thereafter [5] and today an impressive series of enzyme systems is known which depend on nickel as this volume demonstrates. Interestingly, already in 1959 Wacker and Vallee detected nickel as well as other metals in RNAs from diverse biological sources [6]. However, it is uncertain even today if there is a ‘positive’ role for nickel in nucleic acids, though on the ‘negative’ side there is evidence that this metal ion may be responsible for the formation of reactive oxygen species which can then oxidize nucleobases in nucleotides and nucleic acids [7]. This may be one of the reasons why nickel is toxic and carcinogenic as well (see Chapter 17); that nickel may interact with DNA [8] and that it affects gene expression is also known (see Chapter 16). Before dealing with the complexing properties of nucleic acid constituents, it seems appropriate to outline first a few of the basic chemical properties of nickel. Nickel(II) prefers an octahedral coordination geometry [3] and indeed, in all complexes formed with nucleic acid constituents [9] and as discussed in this chapter, Ni2 is hexacoordinate. The transition to square planar complexes, as it occurs with sulfhydryl-containing amino acids [10] and with tri- or higher peptides under deprotonation of amide groups [11] (see also Chapter 3), does not take place with nucleobases or phosphates [9]. The first deprotonation from a metal-ion-bound water molecule on aqueous Ni2 requires quite basic solutions because pKa 10.2 [10], though this value is somewhat misleading because in 1 mM solutions Ni(OH)2 begins to precipitate already near pH 7 [10]. On the other hand, considering the normally low concentrations in biological fluids, Ni2 is commonly sequestered by other ligands, so that hydroxide insolubility is not of concern [10], and also hydroxocomplex formation hardly occurs in the physiological pH range, as is evident from Ni(ATP)(OH)3 which forms with pKa 9.4 [12]. An important distinction between Ni2 and its neighboring M2 ions in the Periodic Table is the relatively slow rate of ligand exchange in and out of the coordination sphere: The first-order water exchange rate constant for Ni2 [log k 4.3 (s1)] is 102 times slower than that for Co2 and 103 times slower than that for Zn2 [10]. Of course, the rate constants for other ligands differ numerically, but the relative ordering remains the same. This relatively slow ligand exchange rate, appropriate for the temperature-jump technique, is the reason why kinetic studies of nucleotide-complex formation have mainly been made with Ni2 [13–20]. These Met. Ions Life Sci. 2, 109–180 (2007)

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data furnished, among others [21–24], evidence that a phosphate-coordinated metal ion may also interact with N7 of the purine residue (see Sections 5.2.3 and 5.3). For a short appraisal of the historical development regarding nucleotide– metal ion interactions refs [25] and [26] should be consulted. In this chapter we focus first on the metal-ion-binding properties, mainly in solution, of the three individual parts a nucleotide is composed of, i.e., the sugar, nucleobase, and phosphate residues. Next we consider complex formation of various nucleotides and discuss their connected isomeric equilibria in aqueous solution. A great deal of the arguments will be based on stability constants; therefore, in many instances the complexes of the neighboring divalent metal ions, especially of Co2, Cu2, and Zn2 will also be considered, to be able to delineate the special properties of the Ni2 systems. In all cases only equilibrium constants were selected that had been determined under conditions where no self-association occurs [24–26], i.e., the monomeric species strongly dominate. Finally, selected examples of Ni2–nucleic acid interactions will be discussed.

2.

NICKEL(II)–SUGAR INTERACTIONS

Knowledge on the binding of metal ions to carbohydrates is scarce and little information exists on Ni2. This is true especially if one concentrates on the ribose or 2-deoxyribose residues (see also Section 2.2), which are of significance for the nucleoside derivatives discussed herein. The reason for this lack of knowledge is that these interactions are weak [27,28] as long as the hydroxyl groups at a ribose ring are not deprotonated. The cis arrangement of the 2- and 3-hydroxyl groups as present in a ribose moiety favors deprotonation of one of the two OH groups because in the resulting anion intramolecular hydrogen bond formation occurs [29]. Yet, this favored deprotonation with pKa 12.5 is far beyond the physiological pH range, meaning that such a deprotonation can occur in a biological system only in a very special environment [30]. However, e.g., it can be facilitated by metal ions like Cu2 which is apparently able to bind to the cis-glycol unit of a ribose moiety in aqueous solution at high pH values [31,32], as proven in experiments with adenosine. In contrast, 2-deoxyadenosine shows no deprotonation of the 3-hydroxyl group under the corresponding conditions.

2.1. Carboxyhydrate Amines: A Strong Primary Binding Site May Help in Hydroxyl Group Coordination In the solid state N-glycosides (glycosylamines) have been studied by X-ray crystallography. The reaction of diamines, such as ethylenediamine (En) or trimethylenediamine (Tn 1,3-diaminopropane), with ketose or aldose sugars allows Met. Ions Life Sci. 2, 109–180 (2007)


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the isolation of compounds which contain the complex units Ni(En)(D-Fru-En)2 [33], Ni(En)(L-Sor-En)2 [34], Ni(L-Rha-Tn)22 [35] or Ni(D-Mal-Tn)22 [36], where Fru fructose, Sor sorbose, Rha rhamnose, and Mal maltose. In the first two examples the octahedral coordination sphere of Ni2 is occupied by the bidentate En and the tetradentate glycosylamine ligand. One nitrogen of En binds to C2 of the sugar unit forming the glycosylamine. This ligand attaches to Ni2 at four sites, i.e., through the 1- and 3-hydroxyl groups and the two N atoms of the En residue. In the latter two examples two tridentate glycosylamines bind to Ni2 via their two N atoms and the (C2)OH group [36]. In a circular dichroism study of Ni2 complexes in solution with N-glycosylamine ligands formed by 1,3-diaminopropane and pentoses such as D -xylose, D -ribose or D -arabinose, it was concluded that two of the tridentate N-glycosylamine ligands coordinate to one Ni2, each ligand being bound in a meridional mode by the primary amino group, the N-glycosidic secondary amino group and the C2 hydroxyl group of the sugar moiety [37]. For the D-ribose derivative this binding mode was confirmed by an X-ray structure analysis [37]. Interestingly, crystalline Ni2 complexes of dianionic glycopyranoside ligands were obtained by reaction of Ni[tris(2-aminoethyl)amine](OH)2 with methyl-Dglucopyranoside or sucrose [38]. In the latter case (C2)O and (C3)O chelation occurs in the glucose part of the disaccharide. It is important to note that here deprotonated hydroxyl groups participate in Ni2 binding. In this context it may also be mentioned that Ni2 complexes of N,N-alkylated ethylenediamine are able to catalyze the C2 epimerization of aldoses and ketoses [39]. Potentiometric and spectroscopic studies in aqueous solution (25C; I 0.15 M, KNO3) indicate that 2-amino-2-deoxy-D-mannose, 2-amino-2-deoxy-D-galactose, and 2-amino-2-deoxy-D-glucose form Ni2 complexes which also involve the hydroxyl groups of the sugars [40]. From these and related studies [41] it becomes evident that the amino group is the major donor towards Ni2 with one of the hydroxyl groups as the second donor which is being deprotonated only at pH 8.

2.2.

Ribose Hydroxyl Groups Are Poor Binding Sites in Complexes of Nucleoside Monophosphates

It is evident that in all the above examples, be it in the solid state or in solution, the amine residue acts as the primary binding site which may facilitate an interaction with a nearby hydroxyl group, though, this is not always the case [42]. In this context one could think that also a nucleobase or a phosphate residue could act as primary binding sites. Therefore it is interesting to consider the solid state structures of several nucleotide-Ni2 complexes, i.e., of [Ni(NMP)(H2O)5] • nH2O species, where NMP2 AMP2 [43], IMP2 [44,45], GMP2 [46] or dGMP [47]. The common feature for this type of structure is the ‘M-N7 base-only’ metal ion bonding with three intramolecular hydrogen bonds, i.e., one between a water Met. Ions Life Sci. 2, 109–180 (2007)

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Figure 1. Molecular structure of the Ni(IMP)(H2O)5 complex which is representative for all [Ni(NMP)(H2O)5]•nH2O species. The intramolecular hydrogen bonds between Ni2-coordinated water molecules and two phosphate oxygens (2.66 and 2.70 Å) as well as the carbonyl oxygen (2.84 Å) are indicated by dotted lines (the distances between the oxygens involved in the hydrogen bonds are given). The structure has been prepared based on the structure coordinates from [44,45], CSD accession code ANIMPH01. For the chemical structure of the nucleobase residue of IMP2, i.e., hypoxanthine, see Figure 3 in Section 3.

ligand and (C6)O or (C6)NH2 and two between water ligands and phosphate oxygens with all NMPs in the anti conformation [48] (Figure 1). A similar structure was obtained with [Ni(3,5-cGMP)(H2O)5] [49], but the nucleotide is here in the syn conformation [48]. Several examples with Ni2 are also known for cis-[M(NMP)2 (amine)x(H2O)y] n-type complexes, where x 02, y 04, and n 02: [Ni(IMP • H)2 (En)(H2O)2] [50], [Ni(GMP • H)2 (En)(H2O)2] [50], [Ni(dGMP)2 (En) 0.7(H2O) 0.6 (H2O)2] 2 [51], and [Ni(GMP)2 (H2O) 4] 2 [51]. The common feature for these structures is the ‘cis-N7-M-N7 base-only’ metal ion bonding with the NMPs in the anti conformation. There are also intramolecular hydrogen bonds, analogous to those observed for the [Ni(NMP)(H2O)5]-type Met. Ions Life Sci. 2, 109–180 (2007)


115

structure [48], which are formed between amine and water ligands and (C6)O and/or phosphate oxygen(s). However, in all these nucleotide complexes no direct hydroxyl-Ni2 bonds and also no intramolecular and only very rarely intermolecular hydrogen bonds between a Ni2-coordinated water molecule and a ribose-hydroxyl group exist, confirming the low affinity of these groups towards Ni2. Evidently, such hydroxyl groups become stronger binding sites only upon deprotonation which may occur in the higher pH range (see also the introductory paragraphs to Section 2).

2.3. A Favorable Steric Setting and a Reduced Solvent Polarity May Promote Metal Ion—Hydroxyl or Carbonyl Group Binding More insight into Ni2 binding to hydroxyl and carbonyl groups in solution encompassing the neutral pH range and having a phosphate group as the primary binding site, can be gained by considering the metal-ion-binding properties of the keto-triose derivative dihydroxyacetone phosphate (DHAP2) and the other two related compounds shown in Figure 2. The combination of coordinating groups seen at C1 and C2 for DHAP2 and glycerol 1-phosphate (G1P2) is representative for many sugar moieties. From a steric point of view, an interaction of a phosphate-coordinated metal ion with the neighboring keto or hydroxyl group is very well possible in both instances (see Figure 2). The questions are: Does such an interaction occur in aqueous solution? Are 7-membered chelates formed as expressed in a simplified way in Equilibrium (1)? C R

C

C

O

O PO

R

O

O

O O PO

C O

M

(1)

O M

For simplicity charges are omitted in Equilibrium (1).

3

3

H2C OH

H2C OH

2

2

C O 1

C O

HC OH 2–

H2C O PO3

DHAP2–

C H3

1

2–

2–

H2C O PO3

O PO 3

G1P2–

AcP2–

Figure 2. Chemical structures of dihydroxyacetone phosphate (DHAP2), glycerol 1-phosphate (G1P2), and acetyl phosphate (AcP2). Met. Ions Life Sci. 2, 109–180 (2007)

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SIGEL and SIGEL

Any kind of chelate formation has to enhance complex stability [52]. Hence, a possibly increased stability of Ni(DHAP) or Ni(G1P), if compared with a pure phosphate coordination, could therefore be attributed to the participation of the oxygen at C2 in Ni2 binding, i.e., Equilibrium (1) would then truly exist and at least in part be on its right side. The experimentally accessible (overall) stability constant as defined by Equation (2), where R-MP2 represents any monophosphate ligand, i.e., in the present case DHAP2 and G1P2, encompasses all isomeric complexes found, namely the ‘open’ (op) and chelated or ‘closed’ (cl) species: Ni2 R-MP2

Ni(R-MP)

(2a)

Ni K Ni(R-MP) [ Ni(R-MP)]([ Ni 2 ][ R-MP 2 ])

(2b)

([ Ni(R-MP)op ][ Ni(R-MP)cl ])([ Ni 2 ][ R-MP 2 ])

(2c)

The stability constant of the open isomer in Equilibrium (1), Ni(R-MP) op, is defined by Equation (3) Ni K Ni(R-MP)op [ Ni(R-MP)op ]([ Ni 2 ][ R-MP 2 ])

(3)

but it is not accessible by a simple direct experimental determination. However, the existence of a linear relationship for families of structurally closely related H ligands between log KM M(L) and pKH(L) is well known [52] and exists also for log Ni H K Ni(R-PO3) versus pKH(R-PO3) plots [25,53], where R-PO32 represents simple monophosphate monoester and phosphonate ligands such as phenyl phosphate, p-nitrophenyl phosphate, methylphosphonate, etc., i.e., R may be any residue which does not affect complex formation (see also Figure 9 in Section 5.2.3). The parameters for the corresponding straight line for Ni(R-PO3) complexes are given in Equation (4): M log K M(L)

H ⴢ pK H(L)

m

(4a)

b

Ni H (0.245 ± 0.023) ⴢ pK H(R-P log K Ni(R-PO O3 ) (0.422 ± 0.147) 3)

(4b)

Evidently, with a known pKHH(R-PO )value, an expected stability constant for the corresponding Ni(R-PO3) complex can be calculated. Equation (4b) is valid in the pKa range 5–8 for aqueous solutions (25C; I 0.1 M, NaNO3), and the error limit (three times the standard deviation) for a calculated log KNi Ni(R-PO ) value is ±0.05 [25,53]. Now a stability difference according to Equation (5) can be defined: 3

3

Ni Ni log DNi/R-MP log K Ni(R-MP) − log K Ni(R-PO 3) Ni Ni log K Ni(R-MP) log K Ni(R-MP)op

Met. Ions Life Sci. 2, 109–180 (2007)

(5a) (5b)


117

The equality of the various terms in Equation (5) is evident. The stability constants KNi Ni(R-MP) for the Ni(DHAP) and Ni(G1P) complexes have been measured and are known [54] as are the acidity constants for the H(DHAP) and H(G1P) H species, pKH(DHAP) 5.90 ± 0.01 and pKHH(G1P) 6.23 ± 0.01, and therefore with Equation (4) the differences defined in Equation (5) can be calculated [55]. The results are: log DNi/DAPH (1.85 ± 0.03) (1.87 ± 0.05) 0.02 ± 0.06 log DNi/G1P

(1.90 ± 0.04) (1.95 ± 0.05) 0.05 ± 0.06

Both log D Ni/R-MP values are zero within the error limits. Hence, no increased complex stability is observed and it must therefore be concluded that Equilibrium (1) is on its left-hand side and that the (C2)O and (C2)OH groups do not participate in Ni2 binding in aqueous solution. Corresponding results have been observed for the complexes of Mn2, Co2, Cu2, and Zn2 [54,55]. However, it needs to be added that a decreased solvent polarity favors weakly coordinating oxygen sites [30]. Indeed, for Cu(DHAP) and Cu(G1P) it has been shown that in water containing 50% 1,4-dioxane (v/v) the chelated species in Equilibrium (1) reach a formation degree of about 45% [54,55]. A similar chelate formation must be anticipated for the corresponding Ni2 complexes in solutions with a reduced dielectric constant or permittivity (f). The importance of the steric orientation for weak interactions follows from the example with acetyl phosphate (AcP2) (Figure 2, on the right). In this case a six-membered chelate may form as is evident from Equilibrium (6) where X O (AcP2) or CH2 (acetonylphosphonate): H3C

X

O

C

P

O

O–

O– M2+

H3C

X

O

C

P

O

O–

O–

(6)

M2+

In fact, for Ni(AcP) a small, but significant stability enhancement is observed (log D Ni/AcP 0.14 ± 0.06) [56] which corresponds to a formation degree of about 30% for the chelate in the intramolecular Equilibrium (6). For the Ni(acetonylphosphonate) system similar results have been obtained [55,56]. It may further be mentioned that the formation degree of the chelated species is hardly affected in mixed ligand complexes as is evidenced from examples with Cu2 [57]. In conclusion, from the information collected in this section it follows that sugar hydroxyl (or carbonyl) groups are weak binding sites which will interact with Ni2 (or other M2) in aqueous solution only under exceptional conditions [30]. Furthermore, from the solid state structures discussed in Section 2.2 for several NMP complexes it follows that N7 of purine-nucleobase residues has clearly a more pronounced affinity for Ni2 ions than a sugar hydroxyl group. Met. Ions Life Sci. 2, 109–180 (2007)

118

3.

SIGEL and SIGEL

INTERACTIONS OF NICKEL(II) WITH NUCLEOBASE RESIDUES

The common purine and pyrimidine nucleobases are shown in Figure 3. We shall concentrate in this section on N9-substituted purines and N3-substituted pyrimidines, the substituent being an alkyl group, and on the nucleosides which carry a (2-deoxy)ribose residue at the corresponding position. Metal ion binding of the free nucleobases is not of relevance in the present context. It is evident from Figure 3 that these nucleobase residues possess quite a number of potential metalion-binding sites [30], yet from a narrow point of view one may say that N7 is the crucial site for purines and N3/(N3) for the pyrimidines. The details will be discussed below.

3.1.

Nickel(II) Complexes of Purine Derivatives

As said, for the three purines depictured in Figure 3 the N7 site dominates the coordination chemistry of these systems. This is even true under rather exceptional circumstances as the following two examples demonstrate: (1) The adeninium cation which is protonated at N1 is coordinated to Ni2 via the N7 site in the complex [Ni(Nta)(adeninium)(H2O)] • 2.5H2O (Nta nitrilotriacetate) as was proven by an X-ray crystal structure study [58]. This means, even protonation at NH2

7

5

9

6

1N

4 3

N

N

6

9

3

9

3

1

O NH

5 6

NH2

N

Gua Guo

3

O

1 NH

R

O

N N

R=

6

N

N

3

5

N

Hyp Ino

NH2

R=H

1NH

R

Ade Ado

6

7

N

N

R

R=H R = ribose

O

O 7

N

1

N

O

3

H3C

NH 6

1

N

O

R Cyt

R

R

Ura

Thy

ribose Cyd

ribose Urd

2'-deoxyribose dThd

Figure 3. Chemical structures of the nucleobases (R H) adenine (Ade), hypoxanthine (Hyp), guanine (Gua), cytosine (Cyt), uracil (Ura), and thymine (Thy), as well as of the nucleosides (R ribose, except for dThd), adenosine (Ado), inosine (Ino), guanosine (Guo), cytidine (Cyd), uridine (Urd), and thymidine (dThd; R 2-deoxyribose). Met. Ions Life Sci. 2, 109–180 (2007)


119

N1 does not prevent metal ion binding at N7 though it certainly diminishes it [59]. (2) Similarly, and also proven by a crystal structure study [60], in the complex [Ni(Tren)(9-ethylguanine – 0.5H)(H2O)] 2 • (ClO4)2.5 • (ClO3) 0.5, where Tren tris(2-aminoethyl)amine, half of the 9-ethylguanine molecules are deprotonated at their (N1)H site, yet in all instances Ni2 coordination occurs via N7. For the three purines of Figure 3 the acid–base properties for several nucleobase derivatives (NB) including their nucleosides have been measured and the stability constants of several Ni2 complexes were determined. The adenine derivatives can accept a proton at N1 and the hypoxanthine or guanine ones at N7. The latter ones can also release a proton from their (N1)H site; hence, the following equilibria hold: H(NB)

NB H

(7a)

H K H(NB) [H ][ NB][H(NB) ]

(7b)

(NB H) H

NB

(8a)

H K NB [(NB H) ][H ][ NB]

(8b)

Note, the expression (NB H) should be read as NB minus H. Of course, both the NB and the (NB H) species are able to form complexes, the stabilities of which are defined by the following equilibria: Ni2 NB

Ni(NB)2

(9a)

M K M(NB) [ Ni(NB)2 ]([ Ni 2 ][ NB])

Ni2 (NB H)

Ni(NB H)

M 2 K M(NB H) [ Ni(NB H) ]([ Ni ][( NB H ) ])

(9b) (10a) (10b)

The results regarding Equilibria (7–10) are summarized in Table 1 for several purine derivatives [61–69]. From the pKa values of the first two entries it follows that the imidazole nitrogens (N7) of all purines are considerably less basic than imidazole itself. This is also true if annelation, which gives 1-methylbenzimidazole (MBI) (pKHH(MBI) 5.67 ± 0.01 [70]), is taken into account. The corresponding observation is made for the pyridine N1 of the adenines. Consequently, all these nucleobase derivatives are present in the neutral pH range in their uncharged form and hence, metal ions may bind without a competition of the proton (Eq. 9). From Table 1, it can be seen that the stability of the Ni(Ado)2 complex is much lower than is expected from the reduced basicity of N1. This is because the (C6)NH2 group leads to a pronounced steric hindrance for metal ion coordination Met. Ions Life Sci. 2, 109–180 (2007)

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SIGEL and SIGEL

Table 1. Negative logarithms of the acidity constants (Equations 7, 8) of some monoprotonated purine nucleobase (NB) derivatives and logarithms of the stability constants (Equations 9, 10) of the corresponding Ni(NB) 2 and Ni(NB – H) complexes as determined by potentiometric pH titration in aqueous solution at 25C and I 0.1 M (NaNO3) together with data for 1-methylimidazole (MIm) and pyridine (Py). a H(NB)

pKa for

(N7)H H(MIm) H(Py) H(Ado) H(9MeA) H(Ino) H(9MeHyp) H(Guo) H(dGuo) H(9MeG) H(9EtG)

7.20 ± 0.02 (2.2 ± 0.2) d (2.96 ± 0.10) f 1.06 ± 0.06 1.87 ± 0.01 2.11 ± 0.04 2.30 ± 0.04 3.11 ± 0.06 3.27 ± 0.03

log KNi Ni(NB) (N1)H

5.34 ± 0.02 3.61 ± 0.03 4.10 ± 0.01 8.76 ± 0.03 9.21 ± 0.01 9.22 ± 0.01 9.24 ± 0.03 9.56 ± 0.02 9.57 ± 0.05

log KNi Ni(NB H)

3.04 ± 0.01 1.94 ± 0.02 0.4 ± 0.2 e 1.15 ± 0.13g

2.8 ± 0.2h

1.53 ± 0.09 1.81 ± 0.06 1.76 ± 0.10

3.20 ± 0.18 3.46 ± 0.07 3.48 ± 0.13

Reference [61] b [62] c [63] c [64] [68] [69] [68] [69] [69] [69]

a

The errors given are three times the standard error of the mean value or the sum of the probable systematic errors, whichever is larger. b In H(MIm) the proton is at N3. c The proton is released from the positively charged (N1)H site. d Estimated micro acidity constant, pkN7-N1 H•N7-N1, for the (N7)H site of Ado under conditions where N1 does not carry a proton [64]. e Value (based on the work of Lönnberg and Arpalahti [65]) taken from the compilation given in [66]. 25C; I 1.0 M, NaClO4. f Micro acidity constant, pkN7-N1 H•N7-N1 for (N7)H of 9MeA [64]. See also footnote (d). g From Ref. [19]. Estimated error limit; 15C, I 1.0 M, NaClO4. h From Ref. [67]. Estimated error limit; 25C, I 1.0 M, NaClO4.

at N1 [62], but also at N7 [71]! Because the steric effect of a carbonyl group is much lower or even not existent [72,73], the stabilities of the Ni(Ino)2 complex and of the Ni(NB)2 species of the last three entries in Table 1 are significantly higher than that of Ni(Ado)2 and this despite the lower basicities of the involved neutral nucleobases. For the Ni(Ado)2 complex a dichotomy exists because metal ion binding may occur either at N1 or N7. In accord with the importance of N7 as a metalion-binding site, as mentioned above, it was twice concluded for Ni(Ado)2 that the N7-coordinated isomer dominates and values of 70% [74] and 76 ± 8% [66] were derived. This agreement is amazing because the assumptions made in the two studies are quite different. It should be noted already here that this dichotomy disappears in nucleoside 5-phosphates due to the guiding effect of the phosphate group(s) (Sections 5.2.3 and 5.3) [25]. Met. Ions Life Sci. 2, 109–180 (2007)


121

As far as Ni(Ino)2, Ni(Guo)2, and related complexes are concerned, the metal ion coordinates in all instances to N7 and commonly also shows a hydrogen bond between a liganded water molecule and the (C6)O group (Figure 1) [60]. Of course, these complexes are also able to be deprotonated at (N1)H and therefore the Equilibria (7) to (10) are connected with each other via the Equilibrium scheme (11) [75]: Ni2+ + NB

Ni Ni(NB)

Ni(NB)2+

H NB

H Ni(NB)

Ni2+ + (NB – H)– + H+

(11)

Ni(NB – H)+ + H+ Ni Ni(NB – H)

This scheme involves four equilibrium constants and because it is of a cyclic nature only three constants are indepent of each other; the size of the fourth constant is automatically determined by the other three as follows from Equations (12) and (13): Ni H Ni H log K Ni(NB) pK Ni(NB) log K Ni(NB H) pK NB

(12)

H H Ni Ni pK Ni(NB) pK NB log K Ni(NB) log K Ni(NB H)

(13)

The acidification of the (N1)H sites in the Ni(NB)2 complexes, as caused by the (N7)-coordinated metal ion, is defined by Equation (14): H H pK a pK NB pK Ni(NB)

(14)

The corresponding results, based on the data given in Table 1, are summarized in Table 2. These results are in accord with the previous conclusion [69] that the acidifying effect of (N7)-coordinated divalent metal ions on the deprotonation of (N1)H sites in guanine (and hypoxanthine) derivatives decreases in the series Cu2 Ni2 Pt2 Pd2, the ∆pKa values being in the order of about 2.2 ± 0.3 1.7 ± 0.15 1.4 ± 0.1 1.4, respectively. These results indicate further that in the (N1)H-deprotonated Ni(NB H) complexes the metal ion is still largely at N7, though some dichotomy involving also (N1) cannot be excluded. The reason why we favor mainly a N7 coordination is the fact that the pKa values for the kinetically inert and (N7)-bonded Pt2 complexes fit perfectly into the above given series. Furthermore, in the already mentioned half-deprotonated [Ni(Tren)(9EtG – 0.5H)(H2O)]1.5 complex Ni2 is always (N7)-bound having in addition a hydrogen bond via a coordinated water molecule to (C6)O [60]. Of course, in the Ni2 complexes of purine-nucleotides, Met. Ions Life Sci. 2, 109–180 (2007)

122

SIGEL and SIGEL

Table 2. Extent of the (N1)H acidification by (N7)-coordinated Ni2 in hypoxanthine and guanine derivatives as defined by Equation (14) (aqueous solution; 25C; I 0.1 M, NaNO3).a Ni(NB) Ni(Ino) 2 Ni(dGuo) 2 Ni(9MeG) 2 Ni(9EtG) 2

pKHNB

log K Ni Ni(NB)

log KNi Ni(NB – H)

pKHNi(NB) b

pKa

8.76 ± 0.03 9.24 ± 0.03 9.56 ± 0.02 9.57 ± 0.05

1.15 ± 0.13 1.53 ± 0.09 1.81 ± 0.06 1.76 ± 0.10

2.8 ± 0.2 3.20 ± 0.18 3.46 ± 0.07 3.48 ± 0.13

7.11 ± 0.24 7.57 ± 0.20 7.91 ± 0.09 7.85 ± 0.17

1.65 ± 0.24 1.67 ± 0.21 1.65 ± 0.10 1.72 ± 0.18

a

For the error limits see footnote a of Table 1; the error limits of the derived data were calculated according to the error propagation after Gauss. b Values for pKHNi(NB)were calculated according to Equation (13) with the values listed above in columns 2–4 and which are taken from Table 1.

even if (N1)H is deprotonated, the phosphate-coordinated metal ion interacts always with N7 [12] because (N1) could only be reached in the syn conformation (see Figure 8 in Section 5.1). This is not achieved because the anti-syn barrier is too high in energy [76]. There are three more points to be added in the present context: (i) The acidification (∆pKa) by Ni2 at N7 is identical within the error limits for all examples in Table 2, even though the acidity of the various (N1)H sites differs significantly. (ii) The acidification of Ni2 observed in the Ni[d(GpG)] complex is with ∆pKa 1.97 ± 0.25 within the error limits the same as given above [77]. (iii) For the (Dien)Pt(9EtG-N7)2 complex it has been shown that the (N7)-coordinated Pt2 not only acidifies (N1)H (∆pKa 1.40 ± 0.06), but that the released proton can be replaced by another M2 ion, such as Mg2 or Cu2, giving complexes of the type (Dien)Pt(N7-9EtG-N1 • M)3 [78]. The same may be surmised for Ni2. This shows that ‘clustering’ of metal ions at a guanine residue is possible; an observation of importance for ribozymes [30].

3.2. Nickel(II) Complexes of Pyrimidine Derivatives Among the three pyrimidine-nucleobase residues shown in Figure 3 only the cytosine moiety is in the physiological pH range not protonated at N3 and hence, freely available for metal ion coordination. Therefore, this residue will be discussed first. It is important to emphasize in this context the ambivalent properties of the cytosine moiety as revealed by crystal structure studies: In the dimeric [(En)Pt(CMP)] 2 complex the square plane around (En)Pt2 is completed by N3 of one CMP2 and a phosphate oxygen of the other [48,79], whereas in Ba(CMP) • 8.5H2O the alkaline earth metal ion is bonded to (C2)O (and the sugar, but not the phosphate) [48,80]. Between these two ‘extremes’ are cases where both N3 and (C2)O participate as, e.g., in the tetrakis(1-methylcytosine)Met. Ions Life Sci. 2, 109–180 (2007)


123

copper(II) perchlorate dihydrate complex [81]. Here, the four N3 atoms of the nucleobases form a CuN4 plane (mean Cu–N distance 2.03 Å) with more weakly bound C2 carbonyls (mean 2.73 Å) above and below this plane. The stability constants of Ni(Cyd)2 and of some related complexes [82] as defined by Equation (9) are listed in column 2 of Table 3. Unfortunately these numbers alone provide little information about the structures of these complexes in solution. M However, if compared with log KM(L) versus pKHH(L) straight-line plots for simple pyridine-type (PyN; open circles) as well as ortho-aminopyridine-type (oPyN; crossed circles) ligands [62] the stability data become revealing as is evident from Figure 4 and several conclusions are immediately reached: (i) The M(oPyN)2 complexes of all metal ions studied [76] are less stable than the M(PyN)2 species; this proves the steric inhibition of an ortho-amino group next to the coordinating pyridine nitrogen. (ii) The data point for the Ni(Cyd)2 complex [82] fits on the reference line defined by the M(oPyN)2 species, meaning that the neighboring carbonyl group does not participate in metal ion binding and that only the steric inhibition of the (C6)NH2 group is in action. (iii) This is different for the Cu(Cyd)2 complex which shows an increased complex stability thus indicating the participation of the (C2)O group in metal ion binding in accord with the mentioned X-ray study [81]. In other words, the steric inhibiting effect of the (C6)NH2 group is partially offset by the (C2)O group. (iv) The stabilities of the Mg(PyN)2 and Mg(oPyN)2 complexes differ only little and in the pH range 3-7 they are independent of the pKa value of the pyridine derivative considered. This indicates [62] that complex formation takes place in an outersphere manner [76]. (v) Furthermore, Mg(Cyd)2 is even more stable than the sterically unhindered Mg(PyN)2 species proving the importance of the (C2)O interaction in M(Cyd)2 complexes for alkaline earth ions [76] as already evidenced above for Ba2 in the solid state [48,80]. To delineate the special properties of Ni2 in M(Cyd)2 complexes further, it is helpful to compare its properties with those of its neighbors in the Periodic Table. For this reason we have calculated the expected stabilities of the M(oPyN)2 complexes of Mg2, Co2, Ni2, Cu2, and Cd2 by using pKHH(Cyd) 4.24 ± 0.02 [82] and Equation (4a) with the straight-line parameters listed in [62]. These calculated stability constants are listed in column 3 of Table 3 and they quantify the stability of the open species in the intramolecular Equilibrium (15): 2 M(Cyd) op

M(Cyd) 2 cl

(15)

Of course, if a M2 interaction, be it innersphere or outersphere, occurs with N3 and (C2)O of Cyd, a ‘closed’ species results. Hence, in M(Cyd) cl2 either fourmembered chelates exist or if a water molecule participates, a six-membered so-called semichelate may form. In addition, a complete outersphere interaction with both sites can also not be excluded. As a consequence, the M(Cyd) cl2 species are actually a mixture of such chelated isomers [76]. The stability enhancement, which must result from chelate formation [52], is defined by the stability Met. Ions Life Sci. 2, 109–180 (2007)

124

SIGEL and SIGEL 1.2

4BrPy

0.8

3MPy

4ClMPy Py Cyd

3ClPy

3,5DMPy

0.4

Mg2+

0.0 0.4

log

M M(L)

4.0

Cu2+

3.0 2.0 1.0 0.0 3.0 2.0

Ni2+

1.0 0.0 1.0

2M5BrPy Tu 2MPy 2A5BrPy 2APy

2.0 2

3

4

5

6

7

8

H

p H(L)

Figure 4. Evidence for the varying coordinating properties of cytidine () depending on H the metal ion involved. This observation is based on the log KM M(L) versus pKH(L) relationship for simple pyridine-type () as well as ortho-aminopyridine-type (⊗) ligands; the reduced stability of the complexes formed with the latter ligands reflects the steric inhibition due to an ortho-amino (or -methyl) group. The least-squares straight reference lines for the simple pyridine-type ligands are defined by the equilibrium constants for the systems containing () (at the top from left to right) 3-chloropyridine (3ClPy), 4-bromopyridine (4BrPy), 4-(chloromethyl)pyridine (4ClMPy), pyridine (Py), β -picoline ( 3-methylpyridine, 3MPy), and 3,5-lutidine ( 3,5-dimethylpyridine, 3,5DMPy) and those for the ortho-aminopyridine-type ligands by the constants for the systems containing (⊗) (at the bottom from left to right) 2-methyl-5-bromopyridine (2M5BrPy), 2-amino-5-bromopyridine (2A5BrPy), tubercidin ( 7-deazaadenosine, Tu), α-picoline ( 2-methylpyridine, 2MPy) and 2-aminopyridine (2APy). All plotted equilibrium constants refer to aqueous solution at 25C and I 0.5 M (NaNO3); the data for Cyd are from [82] and those for the pyridine derivatives from [62].

difference expressed in Equation (16): M M log DM/Cyd log K M(Cyd) log K M(Cyd)op M M log K M(Cyd) log K M( oPyN) log D

Met. Ions Life Sci. 2, 109–180 (2007)

(16a) (16b)


125

M Table 3. Comparison of the measured stability constants, KM(Cyd) (Equations 9, 19), of M M the M(Cyd) 2 complexes with the stability constants, KM(Cyd)op ( KM(oPyN) ; Equation 20), of the isomers with a sole N3 coordination of M2, and extent of the total chelate formation according to Equilibrium (15) in the M(Cyd)2 complexes in aqueous solution at 25C and I 0.5 M (NaNO3) as expressed by KI (Equations 17, 18) and the percentages of M(Cyd) cl2 (Equation 21).a

M2

M log KM(Cyd)

M log KM(Cyd)op

log D M/Cyd

KI

%M(Cyd) 2 cl

Mg2 Co2 Ni2 Cu2 Cd2

0.12 ± 0.04 0.03 ± 0.08 0.14 ± 0.12 1.56 ± 0.06 0.91 ± 0.07

0.06 ± 0.06 0.03 ± 0.08 0.08 ± 0.10 0.79 ± 0.07 0.53 ± 0.09

0.18 ± 0.07 0.00 ± 0.11 0.06 ± 0.16 0.77 ± 0.09 0.38 ± 0.11

0.51 ± 0.24 0.00 ± 0.25 0.15 ± 0.42 4.89 ± 1.22 1.40 ± 0.61

34 ± 11 0 0 83 ± 4 58 ± 11

a

For the error limits see footnotes a of Tables 1 and 2. The values of column 2 are from [82]; all the other values were calculated as described in the text (for details see [76]).

These log D values correspond to the vertical distances in Figure 4 between the experimentally determined points of a given M(Cyd)2 complex (solid circle) and its reference line (crossed circles) and are listed in column 4 of Table 3. With these considerations in mind we may define in a general way for any ligand (L) the intramolecular equilibrium constant, KI, for Equilibrium (15) (L Cyd): K I [ M(L)cl ][ M(L)op ]

(17)

Values for KI may be calculated [52–56,68] according to Equation (18): KI

M K M(L) M K M(L)op

1

(18a)

10 log D 1

(18b)

M M M Equation (16) defines log D whereas KM(L) ( KM(Cyd) ; Eq. 9) and KM(L)op are defined by Equations (19) and (20):

M K M(L)

M K M(L)op

[ M(L)] ([ M(L)op ][ M(L)cl ]) [ M][L] [ M][L]

(19)

[ M(L)op ]

(20)

[ M][L] Met. Ions Life Sci. 2, 109–180 (2007)

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SIGEL and SIGEL

Knowledge of KI allows us now to calculate the formation degree of the closed or chelated species according to Equation (21): %M(L)cl 100ⴢK I (1 K I )

(21)

The results for KI and % M(Cyd) cl2 are listed in columns 5 and 6 of Table 3. Consideration of the percentages of the closed forms of M(Cyd)2 in combination with results from solid state studies is revealing: (i) Most likely the closed complex Mg(Cyd) 2 cl , formed to about 35%, is a semichelate mainly innersphere bound to (C2)O and outersphere bound to N3 (compare also the Ba2 complex discussed above [48,80]). (ii) In the polymeric Cd(dCMP) complex binding of the octahedral Cd2 occurs to both N3 (2.30 Å) and (C2)O (2.64 Å) by formation of a 4-membered ring [83]. The same may be surmised for Cd(Cyd) 2 cl , but in aqueous solution one expects it to be in equilibrium with a semichelate with N3 innersphere and (C2)O outersphere bound. For Cu(Cyd) cl2 the same structures are indicated [76]. (iii) Unfortunately there is no X-ray structure available for a pertinent Ni2 complex. For the closely related Co2, which also does not form significant amounts of a M(Cyd) 2 cl species (see Table 3, column 6), it was shown that in the polymeric Co(CMP) complex the metal ion coordinates to N3 (1.99 Å) and does not interact with (C2)O [84]. One is inclined to assume the same coordination mode for Ni(Cyd)2 and indeed, chelate formation is zero within the error limits (Table 3, column 6). However, such a monodentate coordinated species may actually to some extent be in equilibrium with a further monodentate binding of Ni2 to (C2)O as has been observed in the solid state in an X-ray study of trans-bis[cytosine-(C2)O]bis(ethylenediamine)nickel(II) bis(tetraphenylborate) [85]. Moreover, considering the low slope of the Ni(oPyN)2 reference line seen in Figure 4 (compare with the Mg2 situation) [62], it could well be that a significant amount of the metal ion in Ni(Cyd)2 is also outersphere bound to N3. To conclude, despite the fact that Ni2 does not form significant amounts of a chelate with the N3/(C2)O sites of Cyd, it is evident that its binding to this nucleobase is still of a rather complicated nature. The other two pyrimidine nucleobases, i.e., the uracil and thymine residues, bind strongly to metal ions only after deprotonation of their (N3)H site. This means that carbonyl groups interact significantly with metal ions only if a suitable primary binding site is available (Section 2.3). To establish a sound basis for comparisons, the four Ni2 complexes of (N3)H deprotonated uridine-type ligands (U), 5-fluorouridine, 5-chloro-2-deoxyuridine, uridine, and thymidine ( 2-deoxy-5-methyluridine, see Figure 3) were studied in conjunction with a few related metal ions [86]. The pertinent equilibria are: U

(UH) K UH [( U − H) ][H ][ U]

Met. Ions Life Sci. 2, 109–180 (2007)

(22a) (22b)


M2 (U H)

127

M(U H)

(23a)

M 2 K M(U H) [ M(U H) ]([ M ][( U H) ])

(23b)

H Plots of log KM M(U H) versus pKU result for the four ligand systems in straight lines [86] and these may be compared with the plots discussed above for pyridine- and o-aminopyridine-type ligands. Figure 5 shows the situation for the Ni2 complexes together with the data for the corresponding Mn2 and Cd2 complexes for comparison. From the Mn2 and Cd2 parts of Figure 5 it follows that their M(U H) complexes are more stable than the Mn(PyN) 2 and Cd(PyN) 2 species, whereas the Ni(U H) complexes are less stable than their Ni(PyN) 2 counterparts. Furthermore, the Ni(U H) straight line is placed (although with a somewhat steeper slope) between the lines of the Ni2 complexes of the PyN- and o-PyN-type ligands. This indicates that Ni2 (like Co2 [86]) suffers in its coordination to (N3) of (U H) from a steric hindrance by the neighboring (C2)O/(C4)O groups. This hindrance however, is less pronounced than that by an o-amino (or o-methyl) group. In contrast, in Mn(U H) and Cd(U H) , (C2)O and (C4)O facilitate M2 binding leading thus to an increased stability.

M M(L) or log

log

4.0

2.5

Mn2+ 1.5

Ni2+ (U

M M(U

H)

2.0

H)

PyN 3.5

2.0

3.0 1.5

2.5

1.0 1.0 0.5

(U

H)

PyN oPyN oPyN

0.5

0.0

H)

PyN

1.0 0.5

0.5 2 3 4 5 6 7 8 9 10 11

(U

2.0 1.5

0.5 0.0

Cd2+

oPyN

0.0 2 3 4 5 6 7 8 9 10 11

2 3 4 5 6 7 8 9 10 11

H p H H(L) or p U M H 2 2 Figure 5. Comparison of the log KM(U H) versus pKU relationships () for Mn , Ni , 2 M H and Cd [86] with the corresponding log KM(L) versus pKH(L) relationships [62] for simple pyridine-type (PyN) () and sterically inhibited ortho-amino(methyl)pyridine-type ligands (oPyN) (). For the definition of the data points of the PyN and oPyN systems see legend of Figure 4 (compare from left to right) (25C; I 0.5 M, NaNO3). The straightreference line for the uridinate-type complexes () is defined (from left to right) by 5fluorouridinate, 5-chloro-2-deoxyuridinate, uridinate, and thymidinate (25C; I 0.1 M, NaNO3; the corresponding equilibrium constants are listed in [86]).

Met. Ions Life Sci. 2, 109–180 (2007)

128

SIGEL and SIGEL

In a careful evaluation, taking into account the situation in M(Cyd) 2 complexes [86], it was concluded that the ‘lower limits’ for chelate formation for Mn(U H) and Cd(U H) are 30 and 60%, respectively. In Cd(U H) 4-membered chelates may form, but in aqueous solution it is highly likely that in addition semichelates occur in which Cd2 is innersphere-bound to (N3) and outersphere via water molecules to (C2)O and (C4)O. In contrast, no chelate formation is anticipated for the Ni2 (and Co2) complex of (U H) , i.e., the metal ion coordinates most likely in a monodentate fashion to (N3) of the uridinates [86]. To conclude, the affinity of Ni2 towards the carbonyl group of the pyrimidine nucleobases, be it innersphere or outersphere, is not pronounced.

4.

COMPLEXES OF NICKEL(II) WITH PHOSPHATES

In Figure 6 the general structures for monoesters of mono-, di-, and triphosphates are shown, the symbols employed are R-MP2, R-DP3, and R-TP 4, respectively. It is evident that the most basic phosphate residue will always be the terminal one, which carries a charge of two minus. In addition, the effect of the residue R on the basicity of such a phosphate group will be the more pronounced the closer they are. For example, the effect of a phenyl and a butyl residue in R-MP 2 leads to the pKa values of 5.85 ± 0.01 and 6.72 ± 0.02, respectively [87], whereas the effect of the same residues in R-DP3 leads to the much smaller pKa span of 6.32 ± 0.02 to 6.65 ± 0.02 [88]. In the case of the triphosphates the residue R has practically no effect [89–91]. Pyrimidine-nucleoside O– R

R

O

O

Pα

Pβ

O

O

O–

O–

Pα

O–

O– O

O

O

O

Pγ

O–

R-TP4–

O

Pβ

O–

R-DP3–

O

O– R

O

P

O–

R-MP2–

O

Figure 6. Chemical structures of simple monoesters of monophosphate (R-MP2), diphosphate (R-DP3), and triphosphate (R-TP 4). R represents a residue which does not affect metal ion coordination at the phosphate, i.e., neither in a positive nor negative sense. Met. Ions Life Sci. 2, 109–180 (2007)


129

6.0

M(R-TP) 2

4.0

M(R-DP) 3.0

log

M M(R-P)

5.0

2.0

M(R-MP) 1.0 0.0

2+

Mg

Mn

2+

2+

Fe

2+

Co

Ni

2+

2+

Cu

2+

Zn

Figure 7. Irving–Williams sequence-type plots for the 1:1 complexes of Mg2 through Zn2 formed with mono- (R-MP2), di- (R-DP3), and triphosphate monoesters (R-TP 4) ( R-P). The plotted data are from Table 13 in [94]; these data also represent the stabilities of the M2 complexes of the pyrimidine-nucleoside 5-mono-, di-, or triphosphates (with the exception of Cu(CTP) 2; see [92]) (25C; I 0.1 M, NaNO3).

phosphates furnish representative values for nucleotide comparisons because the pyrimidine residue does commonly not participate in metal ion binding with labile ions like Ni2 [87,88,92]. The corresponding acidity constants for the terminal phosphate groups are pKHH(R-MP) 6.20 for monophosphate monoesters [25,87], pKHH(R-DP) 6.40 for diphosphate monoesters [88], and pKHH(R-TP) 6.50 for triphosphate monoesters [89–92]. The stability constants (defined by analogy to Eq. 2) which correspond to these pKa values [88], are plotted in an Irving–Williams sequence-type fashion for several M(R-MP), M(R-DP) , and M(R-TP) 2 complexes in Figure 7. The figure confirms the well-known observation [93,94] that phosphate complexes do not strictly follow the Irving–Williams series [95]. The complexes of Ni2 are especially unstable; they are always less stable than their counterparts with Mn2 or Zn2. The figure also reveals that addition of a further phosphate 3 unit to R-MP2 (log KNi Ni(R-MP) 1.94 ± 0.05 [94]), giving R-DP , increases the 2 stability of the Ni complexes by approximately 1.6 log units. The addition of one more phosphate unit, giving R-TP 4, has a somewhat smaller effect (see also below), but the stability increase with nearly 1 log unit is still significant. The stability increase of the complexes varies significantly from metal ion to metal ion by going from M(R-MP) to M(R-DP) , i.e., within the relatively large span from 1.6 to 2.4 log units (see Figure 7). In contrast, when going from M(R-DP) to M(R-TP) 2, the stability increase is quite constant within the Met. Ions Life Sci. 2, 109–180 (2007)

130

SIGEL and SIGEL

narrow range of 0.8–1.0 log units (if the special case of Cu2 with its distorted octahedral coordination sphere is ignored). This indicates in our view that outersphere species play a significant role in M(R-MP) complexes (see also Figure 1), which also include six-membered semichelates with one of the terminal oxygens innersphere and the other one outersphere [87]. Such outersphere interactions are hardly of relevance in the corresponding di- and triphosphate species where two neighboring phosphate units allow the formation of six-membered H innersphere chelates. This is in accord with the log KM M(R-MP) versus pKH(R-MP) 2 2 plots where the slopes are much lower (Mg : m 0.208 ± 0.015; Ni : m 0.245 ± 0.023 [53]) (thus indicating partial outersphere binding) than for the H 2 2 log KM M(R-DP) versus pKH(R-DP) plots (Mg : m 0.485 ± 0.119; Ni : m 0.712 ± 0.223 [88]). Clearly, for di- and triphosphates the formation of innersphere complexes is more pronounced compared with monophosphates, owing to the increased negative charge of these ligands. The ratio of innersphere to outersphere species has been estimated for the Ni2 complex of the monoprotonated and thus four-fold negatively charged triphosphate to be 150:1 (from [96] based on data given in [97]). Overall, the situation may be summarized with the earlier statement [96], ‘the lower the charge, the more predominant are outersphere complexes’. How is the situation with phosphate diester groups? These carry only a single negative charge, yet such units are the bridging phosphates of nucleic acids forming their backbone, which means that they are of great biological relevance [30,98,99]. Together with other functional groups, e.g., the N7 sites of purine residues, these P(O)2 units form specific metal ion-binding pockets in large nucleic acids [30,100–103]. However, if an individual phosphate diester bridge is considered as, e.g., in studies of the Zn2/pUpU3 system [104], one has to conclude that the interactions between Ni2 and such a phosphate diester unit ((O)P(O)(O) 2 ) are weak. Zn2 is used here as a Ni2 ‘substitute’ in accord with the so-called Stability Ruler [105] though from Figure 7 it follows that Ni2-phosphate interactions are somewhat weaker than Zn2-phosphate ones. Furthermore, in studies of poly(U) with Mg2 and Ni2 it was found that the apparent stability constants for the two Mg metal ions are identical within the error limits: log KMg(pU)app 1.80 and log Ni KNi(pU)app 1.83 (25C; I 0.10.3 M) [106]. From these results it was concluded ‘that innersphere coordination … does not occur to a significant extent. Rather, the metal ions are bound in these systems mainly by electrostatic forces, forming a mobile cloud’ [106]. Hence, one can deduce that the interaction of Ni2 with phosphate diester groups occurs predominately in an outersphere manner. Indeed, in accord with the low stability, log KNi Ni(H;RibMP) 0.7 [13], a significant part of the Ni2 complex formed with monoprotonated D-ribose 5-monophosphate (RibMP2) exists as an outersphere species. For comparison, the stability of the Ni(RibMP) complex is much larger with log KNi Ni(RibMP) 2.00 ± 0.01 [87]. Met. Ions Life Sci. 2, 109–180 (2007)


5. 5.1.

131

NICKEL(II) COMPLEXES OF NUCLEOTIDES Some General Considerations

Nucleotides exist in solution mainly in the so-called anti conformation. Two such examples are shown in Figure 8 [107,108]. It is the phosphate residue in nucleotides which determines to a very large part the stability of the complexes formed with the biologically important metal ions, including Ni2 [68,109]. It is important to note that this is independent of the kind of nucleobase involved or whether a nucleoside mono-, di-, or triphosphate is considered. For purine-nucleoside 5-phosphates the formation of macrochelates was proposed about 50 years ago [110] and more than 40 years ago it was concluded that NH2 7

– –

O

Pγ O

Pβ

5 6

O

O

O

N

–

–

O

O

Pα

O

N

N 3

O

O

O

4'

1' 3'

ATP 4–

2

4

9

5'

CH2

1

N

2'

O H

O H NH2

– –

O

Pγ O

O

O

O

Pβ O

5 4

–

–

O

O

Pα O

CTP4–

6

O

5'

1

N

CH2

N3 2

O

O

4'

1' 3'

O H

2'

O H

Figure 8. Chemical structures of adenosine 5-triphosphate (ATP 4) and cytidine 5-triphosphate (CTP 4) in their dominating anti conformation [25,48,63,107,108], together with the labeling system of the triphosphate chain; note that the phosphate groups in the NTPs are labeled α , β, and γ, where γ refers to the terminal phosphate group (see also Figure 6). The analogous NMPs and NDPs have the corresponding structures with one or two phosphate groups, respectively. The adenine and cytosine residues in the structures given above for ATP 4 and CTP 4, respectively, may be replaced by one of the other nucleobase residues shown in Figure 3; if this substitution is done in the way the bases are depicted within the plane (Figure 3), then the anti conformation will also result for the corresponding nucleoside 5-phosphates. The abbreviations AMP2, ADP3, ATP 4, IMP2, etc., in this text always represent the 5-derivatives; 2- and 3-derivatives are defined by 2AMP2, 3AMP2, etc.; in a few instances where uncertainties might otherwise occur, the abbreviations 5AMP2, 5ADP3, etc., are also used. Met. Ions Life Sci. 2, 109–180 (2007)

132

SIGEL and SIGEL

they actually exist [21–23,111], i.e., a metal ion coordinated to the phosphate residue of a purine nucleotide may also interact in the dominating anti conformation with N7 of the purine moiety. Only soon thereafter, it was also discovered that macrochelate formation can be inhibited by the addition of, e.g., 2,2-bipyridine (Bpy), and subsequent formation of mixed ligand complexes [12,112,113] (see Section 8.3). Nowadays formation of macrochelates in complexes formed by purine nucleotides with a variety of metal ions including Ni2 is well established [24,109]. It is evident that the formation of such a macrochelate must give rise to the following intramolecular Equilibrium (24): phosphate-ribose-base M2+

I

phosphate-r i M2+ ob s base-e

(24)

Any kind of equilibrium between an ‘open’ isomer, M(L) op, and a chelated or ‘closed’ isomer, M(L) cl, must be reflected in an increased complex stability compared to the situation where only the open complex can form [52]. This stability enhancement can be evaluated as described already in Section 3.2 and defined in Equations (17) to (21), which furnish values for KI (Eqs 17, 18) and thus for the formation degree of the closed species, % M(L) cl (Eq. 21). One important aspect needs to be emphasized and kept in mind when dealing with purine derivatives: All of them show a pronounced tendency for selfassociation, which occurs via stacking of the purine rings [24,94,114]. As a result, experiments aimed at determining the properties of monomeric metal ion complexes of purine nucleosides or their phosphates should not be carried out in concentrations higher than 103 M. To be on the safe side, it is actually recommended that a maximum concentration of only 5 104 M is used (e.g., [53,109]). The equilibrium constants discussed below were mostly obtained by potentiometric pH titrations and the corresponding experimental conditions adhere to the above request.

5.2. 5.2.1.

Complexes of Nucleoside 5-Monophosphates Definition of the Equilibrium Constants

A combination of the information provided in Figures 3 and 8 reveals that there are nucleoside 5-monophosphates (NMP2), which contain a nucleobase that may accept a proton, e.g., AMP2, whereas others contain a nucleobase that can only release one, e.g., UMP2. Therefore, if one neglects a two-fold protonated Met. Ions Life Sci. 2, 109–180 (2007)


133

phosphate group because this is not of relevance for a biological system due to its large acidity (e.g., for H2 (UMP) pKHH2 (UMP) 0.7 ± 0.3 [87]) one has to consider overall the following three deprotonation reactions: H2 (NMP) ±

H(NMP) H

K HH2 ( NMP) [H(NMP) ][H ][H 2 (NMP)± ] H(NMP)

NMP2 H

H K H(NMP) [ NMP 2 ][H ][H(NMP) ]

(NMP )3 H

NMP2 H K NMP

[(NMP H)3− ][H ][ NMP 2 ]

(25a) (25b) (26a) (26b) (27a) (27b)

Based on the structures of the various NMPs, it is clear that Equilibria (25a) and (26a) hold for H2 (CMP) ± and H2 (AMP) ± , and that Equilibria (26a) and (27a) are of relevance for H(UMP) and H(dTMP) , whereas all three Equilibria (25a)–(27a) are needed to describe the acid–base properties of H2 (IMP) ± and H2 (GMP) ± . In all cases Equilibrium (26a) refers to the deprotonation of the P(O)2 (OH) group in the H(NMP) species. Correspondingly, one has to consider the complexes M(H;NMP) and M(NMP), the stabilities of which are defined in Equilibria (28a) and (29a) below: M2 H(NMP)

M(H;NMP)

M K M(H;NMP) [ M(H; NMP) ]([ M 2 ][H(NMP) ])

M2 NMP2

M(NMP)

(28a) (28b) (29a)

M K M(NMP) [ M(NMP)]([ M2 ][ NMP 2 ])

(29b)

At high pH values also Ni(NMP – H) species might form, but at present no information is available about these. M(H;NMP) complexes form with CMP2, AMP2, IMP2, and GMP2. In these cases, M2, including Ni2, is mainly located at the nucleobase residue and the proton at the phosphate group [25,115,116]. However, because the proton is lost with a pKa value of about 5 or below [25,115,116], these species are of relevance for biological systems only under very special conditions. At the common physiological pH of about 7.5 they are not important and therefore not considered further in the present context. Met. Ions Life Sci. 2, 109–180 (2007)

134

5.2.2.

SIGEL and SIGEL

Properties of Pyrimidine-Nucleoside 5-Monophosphate Complexes

At first we shall concentrate on the pyrimidine-NMPs; their acidity constants are listed in Table 4 together with the stability constants of their Ni(NMP) complexes (Eq. 29). In the same table the corresponding data for tubercidin 5-monophosphate (TuMP2 7-deaza-AMP2) are also given. This nucleotide has the structure of AMP2, except that N7 is replaced by a CH unit and this exchange transforms this purine-type nucleotide into one with pyrimidine-type properties (see Sections 5.2.3 and 6). The α-phosphate group is close in distance to the nucleoside residue and its basicity properties are therefore somewhat affected by this residue (see also Section 4). Indeed, the acidity constants of H(NMP) species as defined by Equation (26b) vary roughly between pKHH(NMP) 5.7 and 6.3 [109,117]. As a consequence, a direct comparison of stability constants of M(NMP) complexes, such as those given in Table 4, does not allow unequivocal conclusions regarding the solution structures of these species. M To overcome the indicated handicap, log KM(R-PO (Eq. 29) versus pKHH(R-PO3) 3) 2 (Eq. 26) plots were established for R-PO3 systems (R being a noninteracting residue), i.e., for simple phosphate monoesters, such as phenyl phosphate or n-butyl phosphate, and phosphonate ligands, such as methanephosphonate [53]. The resulting straight lines are defined by Equation (4a). The equilibrium data for the methyl phosphate and the hydrogen phosphate systems fit on these same reference lines, including data for Ni(CH3OPO3) and Ni(HOPO3) [118], as do the data for RibMP2 and for the pyrimidine-NMPs, UMP2 and dTMP, given in Table 4 (see Figure 9 below) [53,87].

Table 4. Acidity constants of protonated pyrimidine-NMPs (Eqs 25–27) and stability constants of their Ni(NMP) complexes (Eq. 29). The corresponding data are also given for H2 (TuMP) ± (aqueous solution; 25C, I 0.1 M, NaNO3).a Acid H2 (CMP) ± H(UMP) H(dTMP) H2 (TuMP) ±

pKHH (NMP)

pKHH(NMP)

pKHNMP

4.33 ± 0.04b

6.19 ± 0.02 6.15 ± 0.01 6.36 ± 0.01 6.32 ± 0.01

9.45 ± 0.02 9.90 ± 0.03d

2

5.28 ± 0.02 e

a

d

log KNi Ni(NMP)

Ref.

1.94 ± 0.06 1.97 ± 0.05 1.92 ± 0.06 2.04 ± 0.08

[87] c [87] [87] [109]

For the error limits see footnotes a of Tables 1 and 2. This proton is released from the (N3)H site (see Figure 3). c See also [115]. d The proton is released from the (N3)H site (see Figure 3). e This proton is released from the (N1)H site of the 7-deaza-purine residue (cf. with the adenine residue in Figure 3). b

Met. Ions Life Sci. 2, 109–180 (2007)


135

In the light of the results discussed in Sections 2.2 and 3.2, it is no surprise that the ribose residue and the uracil and thymine moieties do not participate in metal ion binding in M(RibMP), M(UMP), and M(dTMP) complexes [53]. What is the situation with Ni(CMP)? Application of pKHH(CMP) 6.19 (Table 4) to the straightline parameters summarized for Ni(R-PO3) in Equation (4b) (Section 2.3) allows calculation of the ‘open’ isomer Ni(CMP) op in Equilibrium (24), i.e., log KNi Ni(CMP)op 1.94 ± 0.05. This calculated stability constant is identical with the measured one given in Table 4. Hence, there is no indication for an increased stability of the Ni(CMP) complex, which means that Equilibrium (24) is far on the left-hand side and that no remarkable amounts of macrochelates are formed. At first sight this result for Ni(CMP), which also holds for Cu(CMP) and other M(CMP) species [25], may seem surprising because the ability of the N3 site of the cytosine residue to interact with metal ions is well known (Section 3.2; Table 3). However, it should be recalled that CMP2 exists predominately in the anti conformation (Figure 8) in which N3 is pointing away from the metal ion coordinated at the phosphate group. Evidently, for all divalent metal ions considered (Table 3) the anti–syn energy barrier, which has been estimated to be of the order of about 7 kJ/mol [76], is too large to be overcome by macrochelate formation.

5.2.3.

Properties of Purine-Nucleoside 5-Monophosphate Complexes

Purine-nucleotides represent a favored situation because not only is N7 of the purine moiety able to bind metal ions (Section 3.1; Table 1), but also a metal ion coordinated at the phosphate residue can reach N7 in the dominating anti conformation (Figure 8) as was already pointed out in Section 5.1. Such a possible N7 interaction, depending on the kind of metal ion involved, may give rise to macrochelate formation and if this occurs it must be reflected in an increased complex stability. Indeed, a very significant stability enhancement is observed for the Ni(NMP) complexes of AMP2, IMP2, and GMP2 (Figure 9), whereas that for the corresponding Mg2 complexes is much smaller. This confirms the wellknown high affinity of Ni2 for N sites, the one of Mg2 being much lower [93]. It is evident that the vertical distances between M(NMP) data points and their corresponding straight reference line can easily be quantified. Application of the acidity constant pKHH(NMP) to the straight-line Equation (4b), the parameters of which are listed in [25,53,55], provides the value of the intercept with the straight line and thus the stability constant of the ‘open’ species (Equation 20). In other words, by analogy to Equation (16) one may now define in Equation (30) the stability enhancement, if any, for the M2 complexes of the purine-NMPs: M M log DM NMP log K M(NMP) log K M(NMP)op M M log K M(NMP) log K M(R-PO log D 3)

(30a) (30b)

Met. Ions Life Sci. 2, 109–180 (2007)

136

SIGEL and SIGEL

3.6

GMP2

log

M M(R-PO3 ) or log

M M(NMP)

3.4 3.2

IMP2

3.0 2.8

AMP 2

2.6

PMEA 2

2.4 2.2

PMEA 2

2.0 1.8

Ni2+

1.6

IMP2 AMP 2

GMP

Mg 2+

1.4

EtP 2 MeP 2

UMP 2 dTMP2

1.2

BuP2

1.0 0.8

2

NPhP 2

5.0

PhP2

5.4

5.8

RibMP 2 6.2

6.6

7.0

7.4

7.8

H p H H(R-PO3 ) or p H(NMP)

Figure 9. Evidence for an enhanced stability of the Ni2 and Mg2 1:1 complexes (Eq. 29) of AMP2 (), IMP2 (⊗), and GMP2 (), based on the relationship between H log KM M(R-PO3) and pKH(R-PO3) for M(R-PO3) complexes of some simple phosphate monoester and phosphonate ligands (R-PO32) (): 4-nitrophenyl phosphate (NPhP2), phenyl phosphate (PhP2), uridine 5-monophosphate (UMP2), D -ribose 5-monophosphate (RibMP2), thymidine [ 1-(2-deoxy-β-D -ribofuranosyl)thymine] 5-monophosphate (dTMP2), n-butyl phosphate (BuP2), methanephosphonate (MeP2), and ethanephosphonate (EtP2) (from left to right). The least-squares lines (Equation 4a) are drawn through the corresponding 8 data sets () taken from [87] for the phosphate monoesters and from [53] for the phosphonates. The points due to the equilibrium constants for the M2/NMP systems (), (⊗), () are based on the values listed in Table 5. The equilibrium data for the M2/PMEA systems () are from [53]. These results will be discussed in Section 7.4. The vertical (broken) lines emphasize the stability differences from the reference lines; they equal log D M/NMP and log D M/PMEA, as defined in Equation (30), for the M(NMP) complexes. All the plotted equilibrium constants refer to aqueous solutions at 25C and I 0.1 M (NaNO3). M M Clearly, the expressions log KM(NMP)op and log KM(R-PO are synonymous 3) because the calculated value for M(R-PO3) reflects the stability of the ‘open’ isomer in Equilibrium (24), in which M2 is only phosphate-coordinated. Now, with known values for log DM/NMP ( log D ), the evaluation procedure defined

Met. Ions Life Sci. 2, 109–180 (2007)


137

Table 5. Logarithms of the stability constants of several M(NMP) complexes as determined by potentiometric pH titration, together with the calculated stability constants of the ‘open’ forms, as well as with the enhanced complex stabilities, log D M/NMP (Equation 30), and the extent of intramolecular macrochelate formation according to Equilibrium (24) for aqueous solutions at 25C and I 0.1 M (NaNO3).a,b M(NMP)

M log KM(NMP) (Eq. 29)

M log KM(NMP)op (Eqs 4b,20) c

log D M/NMP (Eq. 30)

KI (Eqs 17,18)

%M(NMP) cl (Eqs 21,24)

Mg(AMP) Mn(AMP) Ni(AMP) Cu(AMP) Zn(AMP)

1.62 ± 0.04 2.23 ± 0.02 2.55 ± 0.04 3.17 ± 0.02 2.38 ± 0.07

1.56 ± 0.03 2.16 ± 0.05 1.94 ± 0.05 2.87 ± 0.06 2.13 ± 0.06

0.06 ± 0.05 0.07 ± 0.05 0.61 ± 0.06 0.30 ± 0.06 0.25 ± 0.09

0.15 ± 0.13 0.17 ± 0.15 3.07 ± 0.60 1.00 ± 0.29 0.78 ± 0.38

13 ± 10 15 ± 11 75 ± 4 50 ± 7 44 ± 12

Mg(IMP) Mn(IMP) Ni(IMP) Cu(IMP) Zn(IMP)

1.68 ± 0.05 2.33 ± 0.04 3.12 ± 0.04 3.51 ± 0.04 2.59 ± 0.04

1.57 ± 0.03 2.16 ± 0.05 1.95 ± 0.05 2.88 ± 0.06 2.13 ± 0.06

0.11 ± 0.06 0.17 ± 0.06 1.17 ± 0.06 0.63 ± 0.07 0.46 ± 0.07

0.29 ± 0.17 0.48 ± 0.22 13.8 ± 2.2 3.27 ± 0.71 1.88 ± 0.48

22 ± 10 32 ± 10 93 ± 1 77 ± 4 65 ± 6

Mg(GMP) Mn(GMP) Ni(GMP) Cu(GMP) Zn(GMP)

1.73 ± 0.03 2.42 ± 0.05 3.50 ± 0.07 3.86 ± 0.04 2.83 ± 0.03

1.57 ± 0.03 2.17 ± 0.05 1.95 ± 0.05 2.89 ± 0.06 2.14 ± 0.06

0.16 ± 0.04 0.25 ± 0.07 1.55 ± 0.09 0.97 ± 0.07 0.69 ± 0.07

0.45 ± 0.14 0.78 ± 0.29 34.5 ± 7.0 8.33 ± 1.55 3.90 ± 0.76

31 ± 44 ± 97 ± 89 ± 80 ±

7 9 1 2 3

a

For the error limits see footnotes a of Tables 1 and 2. The values for AMP are from [116], those for IMP and GMP from [25]). b The acidity constants are for H2 (AMP) ± pKHH2 (AMP) 3.84 ± 0.02 (Eq. 25) [(N1)H site] and pKHH(AMP) 6.21 ± 0.01 (Eq. 26), for H2 (IMP) ± pKHH2 (IMP) 1.30 ± 0.10 (Eq. 25) [mainly (N7)H; see Figure 2 in [68] for the micro acidity constants], pKHH(IMP) 6.22 ± 0.01 (Eq. 26) and pKHIMP 9.02 ± 0.02 (Eq. 27), and for H2 (GMP) ± pKHH2 (GMP) 2.48 ± 0.04 (Eq. 25) [(N7)H site], pKHH(GMP) 6.25 ± 0.02 (Eq. 26) and pKHGMP 9.49 ± 0.02 (Eq. 27) [68]. c The values in this column were calculated with pKHH(NMP) by analogy to Equation (4a) by using the straight-line parameters listed in [25,53,55].

in Equations (17–21) (Section 3.2) can be applied and the formation degree of the macrochelate which appears in Equilibrium (24) can be calculated (Eq. 21). The corresponding results are summarized in Table 5 for the Ni(NMP) complexes of AMP2, IMP2, and GMP2, together with the analogous results of some other metal ions for comparisons. The results of column 6 in Table 5 demonstrate that in all M2 1:1 complexes with purine-NMPs at least some macrochelation occurs. Macrochelation by an Met. Ions Life Sci. 2, 109–180 (2007)

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SIGEL and SIGEL

interaction of the phosphate-coordinated M2 with N7 is thereby proven with the Ni2/TuMP2 system (Table 4). In TuMP2 a CH unit replaces the N7 of AMP2 (Figures 3 and 8) which means that TuMP2 is no longer able to form the macrochelates mentioned. Indeed, the stability constants of its M(TuMP) complexes fit on the reference lines, thus proving that TuMP2 behaves like a simple phosphate monoester that binds metal ions only to its phosphate group [25,109]. Similar to the situation with pyrimidine NMPs and their N3 site, also with purines the N1 site cannot be reached in the anti conformation. The data assembled in Table 5 allow at least two more interesting conclusions: (i) In all instances the formation degree of the macrochelates increases in the order M(AMP) cl M(IMP) cl M(GMP) cl. This increase for M(IMP) cl

M(GMP) cl is in accord with the increased basicity of N7 in GMP2 compared with that of N7 in IMP2 [68]. However, with AMP2 the situation is more complicated: 9-methylguanine has a pKa of 3.11 ± 0.06 [64,119] for its (N7)H site and this value is rather close to the micro acidity constant N7-N1 for the same site in 9-methyladenine, pkH•N7-N1 2.96 ± 0.10 [64]. Hence, the N7 basicity cannot be responsible for the decreased complex stability. Similar to the situation with pyrimidine derivatives as discussed in Section 3.2, the reduced stability by about 0.9 log unit of Ni(AMP), compared with that of Ni(GMP), is to be attributed to the steric inhibition which (C6)NH2 exercises on a metal ion coordinated at N7 [71]. The (C6)O group does not have such an effect. In contrast, it rather promotes the stability by forming outersphere bonds to a water molecule of the N7-bound metal ion [68]. (ii) The stability enhancement for the complexes of all three purine-NMPs is most pronounced with Ni2 and not with Cu2 as one might initially expect based on the Irving–Williams sequence [95]. This observation may be explained by the different coordination geometries of Cu2 and Ni2. The Jahn–Teller-distorted Cu2 has a strong tendency to coordinate donor atoms (especially N) equatorially [11] and thus, three such positions are left at a phosphate-coordinated Cu2, but only the two cis positions are for steric reasons able to form a macrochelate with N7 [109]. In the octahedral coordination sphere of Ni2 five positions are left after phosphate coordination and four of these are sterically accessible for N7 coordination. Hence, Cu2 backbinding to N7 of the purine ring is statistically disfavored by a factor of 12 ( 24) corresponding to 0.3 log unit and indeed, this corresponds to the observed diminished stability enhancement of 0.31 ± 0.08 [ log D Cu/AMP – log D Ni/AMP (0.30 ± 0.06) – (0.61 ± 0.06); Table 5] in comparison with the Ni2 complex. In case of IMP2 and GMP2, the stabilities of the Ni2 and Cu2 complexes differ even more, i.e. 0.54 ± 0.09 [ (0.63 ± 0.07) – (1.17 ± 0.06)] and 0.58 ± 0.11 [ (0.97 ± 0.07) – (1.55 ± 0.09)], but these differences are identical within their error limits. This is expected if the effect is attributed to the (C6)O group. Hydrogen bonding via an apical Met. Ions Life Sci. 2, 109–180 (2007)


139

water molecule of Cu2 to (C6)O should be weak because of the Jahn–Teller distortion. In the case of Ni2, where all bond lengths are similar, such a hydrogen bond formation should be stronger (Figure 1) and indeed, it contributes about 0.25 log unit ( average of 0.54 and 0.58 minus 0.31) to the stability enhancements observed for Ni(IMP) and Ni(GMP). Finally, it needs to be mentioned that in the context of kinetic and calorimetric studies, the complex stabilities have also been investigated of the Ni2 systems with CMP [13,120], AMP [14,20,120–122], IMP [19,122], and GMP [122]. These results agree in general well with the listed equilibrium constants, especially if the differences in the experimental conditions are taken into account. The most remarkable observation in these studies is that under certain conditions Ni(AMP)22 complexes are also observed [14,120–122] and their formation may be attributed to stacking between the purine residues in these 1:2 species [122]. This interpretation is in accord with the fact that no Ni(CMP)22 complexes were observed [13,120]. Indeed, pyrimidines stack much less than purines [24,94,114]. However, Ni(IMP)22 and Ni(GMP)22 species were also found [122] and their thermodynamic parameters (DH) were measured. It was concluded that the stacking interaction decreases in the series Ni(AMP)22 Ni(GMP)22 Ni(IMP)22 [122] and this order reflects that of the self-association, adenosine guanosine inosine, which also occurs via stacking [24,114].

5.3. Complexes of Nucleoside 5-Di- and Triphosphates All the NDPs and NTPs considered here (Figures 3 and 8) form Ni(H;NDP) and Ni(H;NTP) complexes [88,92,116,123,124], the proton being mainly at the terminal phosphate group. For the reasons already indicated in Section 5.2.1 for the M(H;NMP) species, the corresponding Ni(H;NDP) and Ni(H;NTP) complexes are also expected to play no, or at best only a minor role at physiological pH and will therefore not be discussed here. In the NDP3 species the residue R (Figure 6) is still close enough to the terminal β -phosphate group to have a small influence on the basicity of this group. M Therefore log KM(R-DP) versus pKHH(R-DP) straight-line plots were constructed [88] to detect any possible stability enhancements for the complexes of purine-nucleoside 5-diphosphates. The important issue at this point is that in all instances the data points of the pyrimidine-nucleoside 5-diphosphate complexes, including Ni(UDP) , Ni(dTDP) , and Ni(CDP) , fall within the error limits on the reference line constructed from ligands such as phenyl diphosphate or n-butyl diphosphate, as can be seen in Figure 10. Hence, in none of these cases does the pyrimidine-nucleobase interact with the metal ion coordinated at the phosphate residue. This corresponds to the observations discussed in Section 5.2.2 for the complexes of the pyrimidine-NMPs. Met. Ions Life Sci. 2, 109–180 (2007)

140

SIGEL and SIGEL 6.0

GDP 3 5.8

IDP3

5.6

Cu2+

ADP3

5.4

M M(NDP)

5.2 5.0

log

M M(R-DP)

or log

4.8 4.6

GDP 3

4.4

IDP3 4.2 4.0

ADP3

3.8

Ni2+

3.6 3.4 3.2

PhDP3 MeDP3

6.2

p

BuDP3

dTDP3 CDP3

UDP3

6.4 H H(R-DP)

6.8

6.6

or p

H H(NDP)

Figure 10. Evidence for an enhanced stability of the Ni2 and Cu2 1:1 complexes (defined in analogy to Equation 29) of ADP3 (), IDP3 (⊗), and GDP3 (), based on M the relationship between log KM(R-DP) and pKHH(R-DP) for simple M(R-DP) complexes (), 3 where R-DP phenyl diphosphate (PhDP3), methyl diphosphate (MeDP3; not measured with Cu2), uridine 5-diphosphate (UDP3), cytidine 5-diphosphate (CDP3), thymidine [ 1-(2-deoxy-β -D -ribofuranosyl)thymine] 5-diphosphate (dTDP3), and n-butyl diphosphate (BuDP3) (from left to right). The least-squares lines (Equation 4a) are drawn through the six indicated (in the case of Cu2 five) data sets taken from [88]. The points due to the M2NDP systems are based on the values listed in Table 6. The vertical dotted or broken lines emphasize the stability differences from the reference lines; they equal log D M/NDP as defined by analogy to Equation (30) (see also Table 6, column 4). All the plotted equilibrium constants refer to aqueous solutions at 25C and I 0.1 M (NaNO3). Met. Ions Life Sci. 2, 109–180 (2007)


141

The situation for the complexes of the pyrimidine-nucleoside 5-triphosphates is also quite similar. Here all the H(NTP) 3 species have the same acidity constant, i.e., pKHH(NTP) 6.50 ± 0.05 [89–92] (see also Section 4 and footnote b of Table 7). Indeed, for a given metal ion the stability constants of the M(UTP) 2, M(dTTP) 2, and M(CTP) 2 complexes are identical within the error limits, with the single exception of Cu(CTP) 2 [92]. Thus, these values can be averaged to obtain the stability of the ‘open’ form in Equilibrium (24), i.e., of the complex in which M2 is coordinated only to the phosphate chain [92,124]. With the above information at hand we are now in the position to investigate the situation for the complexes of purine-NDPs and purine-NTPs by analogy to the procedure discussed in Section 5.2.3 for M(NMP) complexes. Consequently, all the equations applied earlier, i.e., Equations (17) to (21) and (25) to (30), are also valid here as long as L or NMP2 are replaced by NDP3 or NTP 4. It follows from the enhanced stabilities of the purine-NDP complexes seen in Figure 10 that the formation degree of the macrochelated species according to Equilibrium (24) increases within the series Ni(ADP) Ni(IDP)

Ni(GDP) . Indeed, this is confirmed by the numbers of % M(NDP)cl as listed in column 6 of Table 6. Exactly the corresponding observation is made for the Ni(NTP)2 complexes, the data of which are listed in Table 7. At this point it is interesting to compare the results assembled in Tables 5 to 7. Of the many comparisons possible, only a few are to be indicated here. The extent of macrochelate formation in the Ni2 complexes of adenine-nucleotides follows the series Ni(AMP) cl Ni(ADP)cl Ni(ATP)2. This means that the steric fit for a N7 interaction is most ideal in the Ni(AMP) complex despite its lowest overall stability (see also Section 5.2.3). This is confirmed by the orders of Ni(AMP) cl Cu(AMP) cl, Ni(ADP)cl Cu(ADP)cl, and Ni(ATP) cl2 Cu(ATP) cl2; consequently, also Cu(AMP) cl Cu(ADP)cl Cu(ATP) cl2 holds. For situations with a lower formation degree of macrochelates, e.g., with Mg2 or Mn2, M(AMP) cl M(ADP)cl M(ATP) cl2 holds. For all complexes, including those with Ni2, and for all purine nucleotides the extent of macrochelate formation increases for the nucleobase residues adenine hypoxanthine guanine. This order is due to the steric influence of the (C6)NH2 group in adenine, the likely partial (outersphere) participation of the (C6)O unit, and the higher basicity of N7 in guanine than in hypoxanthine as discussed already for the NMPs in Section 5.2.3. The significant formation degrees of the macrochelates involving N7, of the order of 69 and 88% for Ni(ITP) cl2 and Ni(GTP) cl2, respectively (Table 7), leads to an acidification of the corresponding (N1)H sites. This acidification amounts to ∆pKa 0.87 and 1.15 for Ni(ITP)2 and Ni(GTP)2, respectively [12]. Consequently, the addition of 2,2-bipyridine leading to Ni(Bpy)(NTP)2 complexes with intramolecular stacks (Section 8.3) between the pyridyl and purine residues [12,114], diminishes the acidification considerably because of the removal of the metal ion from N7 [12]. Met. Ions Life Sci. 2, 109–180 (2007)

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Table 6. Comparison of the measured stability constants, KM M(NDP), of the M(ADP) , M(IDP) and M(GDP) complexes with the stability constants, KM , of the corM(NDP)op responding isomers with a sole diphosphate coordination of M2, and extent of the intramolecular macrochelate formation according to Equilibrium (24) in the mentioned M(NDP) complexes in aqueous solution at 25C and I 0.1 M (NaNO3).a,b

M(NDP)

log KM M(NDP) (Eq. 29) c

log KM M(NDP)op (Eqs 4a,20) c,d

log D M/NDP (Eq. 30) c

KI (Eqs 17,18) c

%M(NDP)cl (Eqs 21,24) c

Mg(ADP) Mn(ADP) Ni(ADP) Cu(ADP) Zn(ADP)

3.36 ± 0.03 4.22 ± 0.02 3.93 ± 0.02 5.61 ± 0.03 4.28 ± 0.05

3.30 ± 0.03 4.12 ± 0.03 3.54 ± 0.06 5.27 ± 0.04 4.12 ± 0.03

0.06 ± 0.04 0.10 ± 0.04 0.39 ± 0.06 0.34 ± 0.05 0.16 ± 0.06

0.15 ± 0.11 0.26 ± 0.10 1.45 ± 0.36 1.19 ± 0.25 0.44 ± 0.19

13 ± 9 21 ± 7 59 ± 6 54 ± 5 31 ± 9

Mg(IDP) Mn(IDP) Ni(IDP) Cu(IDP) Zn(IDP)

3.33 ± 0.03 4.22 ± 0.04 4.27 ± 0.05 5.69 ± 0.10 4.34 ± 0.05

3.29 ± 0.03 4.11 ± 0.03 3.52 ± 0.06 5.25 ± 0.04 4.09 ± 0.03

0.04 ± 0.04 0.11 ± 0.05 0.75 ± 0.08 0.44 ± 0.11 0.25 ± 0.06

0.10 ± 0.11 0.29 ± 0.15 4.62 ± 1.01 1.75 ± 0.68 0.78 ± 0.24

9±9 22 ± 9 82 ± 3 64 ± 9 44 ± 8

Mg(GDP) Mn(GDP) Ni(GDP) Cu(GDP) Zn(GDP)

3.39 ± 0.04 4.35 ± 0.06 4.51 ± 0.03 5.85 ± 0.04 4.52 ± 0.03

3.29 ± 0.03 4.11 ± 0.03 3.52 ± 0.06 5.25 ± 0.04 4.09 ± 0.03

0.10 ± 0.05 0.24 ± 0.07 0.99 ± 0.07 0.60 ± 0.06 0.43 ± 0.04

0.26 ± 0.14 0.74 ± 0.27 8.77 ± 1.51 2.98 ± 0.52 1.69 ± 0.26

21 ± 9 42 ± 9 90 ± 2 75 ± 3 63 ± 4

a

For the error limits see footnotes a of Tables 1 and 2. The values for ADP are from [116], those for IDP and GDP from [123]). b The acidity constants [116,123] are for H2 (ADP) pKHH2 (ADP) 3.92 ± 0.02 (Eq. 25) c [(N1)H site] and pKHH(ADP) 6.40 ± 0.01 (Eq. 26),c for H2 (IDP) pKHH2 (IDP) 1.82 ± 0.03 (Eq. 25) c [mainly (N7)H], pKHH(IDP) 6.38 ± 0.02 (Eq. 26) c and pKH(IDP) 9.07 ± 0.02 (Eq. 27),c and for H2 (GDP) pKHH2 (GDP) 2.67 ± 0.02 (Eq. 25) c [(N7)H site], pKHH(GDP) 6.38 ± 0.01 (Eq. 26) c and pKHH(GDP) 9.56 ± 0.03 (Eq. 27) c. c The equation numbers refer to analogous situations, i.e., L or NMP2 need to be replaced by NDP3. d The values in this column were calculated with pKHH(NDP) by analogy to Equation (4a) by using the straight-line parameters listed in [88].

There is one more point in the context of macrochelate formation. N7 may be innersphere, but also outersphere bound to the already phosphate-coordinated metal ion. For example, based on several methods it was concluded for Ni(ATP) 2that about 30% are N7 innersphere, 25% are N7 outersphere, and 2 45% exist as Ni(ATP) op , i.e., the open form in Equilibrium (24) [25,89]. In contrast, macrochelation occurs with Mg2 only outersphere at N7 and with Cu2 only innersphere [89]. Based on spectrophotometric measurements, it was Met. Ions Life Sci. 2, 109–180 (2007)


143

2 Table 7. Comparison of the measured stability constants, KM M(NTP), of the M(ATP) , M(ITP) 2 and M(GTP) 2 complexes with the stability constants, KM , of the corM(NTP)op responding isomers with a sole coordination of M2 to the triphosphate chain, and extent of the intramolecular macrochelate formation according to Equilibrium (24) in the mentioned M(NTP) 2 complexes in aqueous solution at 25C and I 0.1 M (NaNO3).a,b

M(NTP) 2

log KM M(NTP) (Eq. 29) c

log KM M(NTP)op (Eq. 20) c,d

log D M/NTP (Eq. 30) c

KI (Eqs 17,18) c

%M(NTP)cl (Eqs 21,24) c

Mg(ATP) 2 Mn(ATP) 2 Ni(ATP) 2 Cu(ATP) 2 Zn(ATP) 2

4.29 ± 0.03 5.01 ± 0.08 4.86 ± 0.05 6.34 ± 0.03 5.16 ± 0.06

4.21 ± 0.04 4.93 ± 0.03 4.50 ± 0.03 5.86 ± 0.03 5.02 ± 0.02

0.08 ± 0.05 0.08 ± 0.08 0.36 ± 0.06 0.48 ± 0.04 0.14 ± 0.06

0.20 ± 0.14 0.20 ± 0.22 1.29 ± 0.32 2.02 ± 0.28 0.38 ± 0.19

17 ± 10 17 ± 15 56 ± 6 67 ± 3 28 ± 10

Mg(ITP) 2 Mn(ITP) 2 Ni(ITP) 2 Cu(ITP) 2 Zn(ITP) 2

4.29 ± 0.04 5.21 ± 0.06 5.01 ± 0.10 6.71 ± 0.10 5.32 ± 0.06

4.21 ± 0.04 4.93 ± 0.03 4.50 ± 0.03 5.86 ± 0.03 5.02 ± 0.02

0.08 ± 0.06 0.28 ± 0.07 0.51 ± 0.10 0.85 ± 0.10 0.30 ± 0.06

0.20 ± 0.17 0.91 ± 0.31 2.24 ± 0.75 6.08 ± 1.63 1.00 ± 0.28

17 ± 11 48 ± 8 69 ± 7 86 ± 3 50 ± 7

Mg(GTP) 2 Mn(GTP) 2 Ni(GTP) 2 Cu(GTP) 2 Zn(GTP) 2

4.31 ± 0.04 5.36 ± 0.03 5.42 ± 0.04 7.38 ± 0.08 5.52 ± 0.05

4.21 ± 0.04 4.93 ± 0.03 4.50 ± 0.03 5.86 ± 0.03 5.02 ± 0.02

0.10 ± 0.06 0.43 ± 0.04 0.92 ± 0.05 1.52 ± 0.08 0.50 ± 0.05

0.26 ± 0.17 1.69 ± 0.25 7.32 ± 0.96 32.11 ± 6.10 2.16 ± 0.36

21 ± 11 63 ± 3 88 ± 1 97 ± 1 68 ± 4

a

For the error limits see footnotes a of Tables 1 and 2. The values for ATP are from [92,124], those for ITP and GTP from [124]). b The acidity constants [90] are for H2 (ATP) 2 pKHH2 (ATP) 4.00 ± 0.01 (Eq. 25) c [(N1)H site] and H 6.47 ± 0.01 (Eq. 26),c for H2 (ITP) 2 pKHH2 (ITP) 2.19 ± 0.05 (Eq. 25) c [mainly (N7)H; pKH(ATP) see [90]], pKHH(ITP) 6.47 ± 0.02 (Eq. 26) c and pKHITP 9.11 ± 0.03 (Eq. 27),c and for H2 (GTP) 2 H pKHH2 (GTP) 2.94 ± 0.02 (Eq. 25) c [(N7)H site], pKH(GTP) 6.50 ± 0.02 (Eq. 26) c and pKHGTP c 9.57 ± 0.02 (Eq. 27). c The given equation numbers refer to analogous situations, i.e., L or NMP2 need to be replaced by NTP 4. d M 4 log KM pyrimidine-nucleoside 5-triphosphate; this M(NTP)op KM(PyNTP), where PyNTP means, for each metal ion the stability constants of the M(UTP) 2, M(dTTP) 2, and M(CTP) 2 complexes have been averaged, with the exception for Cu2, where only the values for Cu(UTP) 2 and Cu(dTTP) 2 have been used [92] (see also [124]).

suggested for Ni(ADP) that about 20% occur as Ni(ADP) op and that from the remaining 80% of Ni(ADP)cl about 65% exist as N7 innersphere and 15% as N7 outersphere macrochelated species [125]. For the Ni2 complexes of IDP3 or ITP 4 and GDP3 or GTP 4 to the best of our knowledge no information exists.

Met. Ions Life Sci. 2, 109–180 (2007)

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SIGEL and SIGEL

It is well known that metal ions promote the dephosphorylation of NTPs [26]. However, Ni2 is one of the least effective divalent metal ions; e.g., for the promoted hydrolysis of ATP at pH 7.5 one observes the order Cu2 Cd2 Zn2 Mn2 Ni2 Mg2 [26]. This ineffectiveness may be due to the slow rate of exchange of ligands out of the Ni2 coordination sphere [126] and/or to an especially ‘regular’ α,β,γ -coordination of this hexacoordinate metal ion to the triphosphate chain. The latter would render the formation of α,β -coordinated Ni2 more difficult compared, e.g., with Cu2 or Zn2, and therefore of course also the binding of the second metal ion to the γ -phosphate group [26]. Note, M2 (NTP) complexes are especially reactive in the hydrolysis reaction, which is actually a transphosphorylation, if they exist in the M(α,β)-M(γ) coordination mode because this facilitates the break between the β - and γ -phosphate groups [26,94]. It has recently been pointed out that trivalent nickel in (cyclam)Ni3, cyclam 1,4,8,11-tetraazacyclotetradecane, is stabilized by P2O74 and ATP 4 [127]. This observation is in accord with a similar one where (cyclam)Ni2 and other (N4-ligand)Ni2 complexes were oxidized by KHSO5 to Ni3 species which then interacted with nucleotides to various degrees [128]. In fact, it is well known that phosphate ligands stabilize high oxidation states, as has also been observed for Mn3 [129]. Furthermore, oxidation of Mn2 to Mn3 in the presence of ATP 4 at a pH of about 8 promotes the dephosphorylation reaction of this triphosphate dramatically [26,130]. These types of redox-triggered hydrolysis reactions might be of importance for signal transmission in biological systems [91,131].

6.

COMPLEXES OF SOME LESS COMMON NUCLEOTIDES

Tubercidin 5-monophosphate is synthesized by molds and fungi. This nucleotide is identical in its structure with AMP (Figures 3 and 8) except that N7 is substituted by CH [25,109]. As already discussed, TuMP2 behaves in its binding properties towards Ni2 like a simple phosphate monoester ligand because N7 is not present (Table 4; Section 5.2.3). The N1 site cannot be reached by a phosphatecoordinated metal ion and thus, its presence in TuMP2 has no effect. It may be added here that the steric inhibition exercised by the (C6)NH2 group on Ni2coordination to N7 has been quantified with 9-methyl-1,3-dideazaadenine, also known as 1-methyl-4-aminobenzimidazole [71]. This is mentioned here because this compound and derivatives thereof are often used as structural analogs of their corresponding parent adenine compounds.

6.1.

Complexes of 2- and 3-Nucleoside Monophosphates

Two other adenine-nucleotides, the Ni2 complexes of which have been studied, are 2AMP2 and 3AMP2 [117,132]. What types of macrochelates are possible Met. Ions Life Sci. 2, 109–180 (2007)


145

with these two AMPs? By considering their structures (cf. Figure 8) it is evident that N7, though crucial for the properties of the M(5AMP) complexes (Section 5.2.3; Table 5), is for steric reasons not accessible to a metal ion already bound to either the 2- or 3-phosphate group. Indeed, the stability enhancements observed for Ni(2AMP) and Ni(3AMP) are with log D Ni/2AMP log D Ni/3AMP 0.06 ± 0.07 (3σ) (defined by analogy to Equation 30) very small [117] (cf. log D Ni/5AMP 0.61 ± 0.06; Table 5) and within the error limits also identical allowing no sophisticated conclusions. Nevertheless, one might be tempted to postulate some chelate formation with the neighboring OH groups of the ribose ring for both AMPs. However, in the light of the discussion in Sections 2.2 and 2.3 the above indication appears as highly unlikely and, based on the results obtained with Cu(2AMP) and Cu(3AMP), it can actually be excluded. The steric conditions for 2AMP2 and 3AMP2 to form such seven-membered chelates are identical and therefore equivalent properties for both complexes are expected, yet the results are log D Cu/2AMP 0.26 ± 0.08 and log D Cu/3AMP 0.08 ± 0.08 which corresponds to formation degrees of 45 ± 10% and 17 ± 16% for Cu(2AMP) cl and Cu(3AMP) cl, respectively [117]. Hence, different structural qualities of the two AMPs must be responsible for the different properties of the two complexes. The obvious conclusion is that in Cu(2AMP) partial macrochelate formation occurs by an interaction of the 2-phosphate-coordinated metal ion with N3 of the adenine residue. Actually, 2AMP2 in its preferred anti conformation is perfectly suited for this type of macrochelate formation [117], whereas in Cu(3AMP) an N3 interaction can only be achieved if the nucleotide adopts the less favored syn conformation – and this costs energy and results thus in the observed lower macrochelate formation degree. The following results obtained for the Ni2 complexes of 2GMP2, 3GMP2, and 2-deoxy-3GMP2 [133] corroborate the above conclusions: log DNi 2 ′ GMP

0.70 ± 0.06

%Ni(2 ′ GMP)cl

80 ± 3

log DNi 3′ GMP

0.56 ± 0.11

%Ni(3′ GMP)cl

72 ± 8

log DNi 2 ′ d 3′ GMP 0.55 ± 0.08

%Ni(2 ′ d3′ GMP)cl 72 ± 5

The stability enhancement for Ni(2GMP) is somewhat larger than that of Ni(3GMP), being a reflection of the anti/syn transformation. The most important conclusion from this list is the fact that removal of the 2-hydroxy group from the ribose ring does not affect the stability enhancement and the degree of chelate formation. Hence, one may conclude that the 2-OH group is not involved in metal ion binding. It should be added here that it is becoming increasingly clear from solution [134,135] as well as X-ray crystal structure studies that the N3 site of a purine may bind metal ions [136-141]. For example, Pt2 binds via N3 to guanine [136] and adenine [137] derivatives, Pd2 to N6,N6,N9-trimethyladenine Met. Ions Life Sci. 2, 109–180 (2007)

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SIGEL and SIGEL

[138], Rh to 8-azaadenine [139] as well as Cu2 [140] and Ni2 [141] to neutral adenine.

6.2. Complexes of Orotidinate 5-Monophosphate In Figure 11 three more nucleotides are shown [142–144] for which stability constants of their Ni2 complexes have been measured. Orotidinate 5-monophosphate (OMP3) (Figure 11, top) is involved in the biosynthesis of pyrimidinenucleotides: OMP3 is decarboxylated to UMP2 (cf. Figures 3 and 8), which is then further transformed e.g., into UTP 4 or CTP 4 (Figure 8). OMP3 itself exists predominately in the depictured syn conformation (i.e., the (C2)O group being above the ribose ring). The carboxylic acid group at C6 of OMP is very acidic and gets deprotonated with pKa 1.46 ± 0.10 [145]. Due to the resulting negative charge the deprotonation of the P(O)2 (OH) group of OMP occurs at a slightly higher pH, i.e., pKa 6.40 ± 0.02 [145], than that of the corresponding group of UMP, i.e., pKHH(UMP) 6.15 ± 0.01 (Table 4). Consequently, at the physiological pH of 7.5 the species OMP3 and UMP2 dominate. For Ni(UMP) we have seen that the uracil moiety does not participate in metal ion binding (Section 5.2.2) and that its stability constant fits on the reference line (Figure 9). This is different for the stability of Ni(OMP) which is by log D Ni/OMP 0.36 ± 0.07 above this line and hence, this complex is more stable than expected on the basis of the basicity of the phosphate group. This enhanced complex stability is solely a charge effect of the non-coordinating (C6)COO group as follows from the average of the stability enhancements observed for 10 different M2 ions in their M(OMP) complexes, i.e., log DM/OMP 0.40 ± 0.06 (3σ) (defined by analogy to Eq. 30) [145]. At higher pH values when the (N3)H site is becoming deprotonated, the different metal ions behave differently: now Ni(OMP – H)2 exists as a macrochelate with a formation degree of 74 ± 7% (see Equilibrium 24); the phosphate-bound Ni2 interacts with the (N3) site [145], which is sterically easily reached (see Figure 11, top).

6.3. Complexes of Xanthosinate 5-Monophosphate Just as OMP is an intermediate in the metabolism of pyrimidines, XMP is one in the metabolism of purines. It is most important to note that XMP exists at the physiological pH of 7.5 as a xanthine-deprotonated (XMP H)3 species, more exactly written as (X – H • MP)3 (see Figure 11) [142]. This fact has so far mostly been overlooked [142,146] and XMP is commonly shown in textbooks by analogy to GMP2 (see Figures 3 and 8), which is not correct. It must further be noted that in XMP the deprotonation of the xanthine moiety (pKa 5.30 ± 0.02) takes place

Met. Ions Life Sci. 2, 109–180 (2007)


147

O 3

– –

4

HN O

O

P

2

O

5'

CH2

O

1

O

N

O

4'

1' 2'

3'

O H

5 6

COO–

OMP3–

O H

O

...

O

7

N –

O

O O

P

4 3

9

5'

CH2

O

5

4'

N O

.... 1

N H

1'

O H

O H

N

N

...

O

5

9

N

1

6

4 3

NH

N

...–. ..O

(XMP – H)3–

2'

3'

7

–

....

–

6

= (X – H·MP)3–

5'

2–

H2C O PO3 4'

HC OH

2–

H2C O PO3 HC OH

3'

HC OH

H2C OH

2'

HC OH

G1P2–

1'

H2C H 3C 8 7

H3C

9 6

N

10 5

N

N 1

4

O 2 3

NH

O

FMN2– Figure 11. Chemical structures of orotidinate 5-monophosphate (OMP3), xanthosine 5-monophosphate (XMP2), which actually exists at the physiological pH of about 7.5 as xanthosinate 5-monophosphate, (XMP – H)3, with a tautomeric equilibrium between the (N1) (N3)H and (N1)H/(N3) sites [142,143], and flavin mononucleotide (FMN2 riboflavin 5-phosphate) as well as for comparison of glycerol 1-phosphate (G1P2). OMP3 is shown in its dominating syn conformation [144] and (XMP – H)3 in its dominating anti conformation [48,63,107].

Met. Ions Life Sci. 2, 109–180 (2007)

148

SIGEL and SIGEL

before that of the P(O)2 (OH) group (pKa 6.45 ± 0.02) [142]. Naturally, these acid–base properties affect the coordination chemistry of XMP [143]. The M(XMP) complexes are quite special and they are better written as (M • X – H • MP • H) ± to indicate that the phosphate group carries a proton, and that the metal ion is coordinated to N7, possibly also undergoing an (outersphere) interaction with (C6)O. With the nine metal ions Ba2, Sr2, Ca2, Mg2, Mn2, Co2, Cu2, Zn2, and Cd2 the average stability enhancement of these complexes amounts to log DM/XMP 0.46 ± 0.11 (3σ) [143]. These stability increases are thereby based on the basicity- and charge-corrected stabilities of the M(xanthosinate) complexes [143]. This equality indicates that the interaction of the N7-bound M2 ions with the monoprotonated phosphate group occurs in an outersphere manner. The stability enhancement for the (Ni • X – H • MP • H) ± complex is somewhat smaller, log DNi/XMP 0.20 ± 0.18, and appears to reflect the notoriously ‘low’ affinity of Ni2 towards phosphate groups (see Section 4 and Figure 7). A further explanation could be that Ni2 is partly coordinated to the (N1) site [74] and then the P(O)2 (OH) group could not be reached for macrochelate formation. The observed stability enhancement for (Ni • X – H • MP • H) ± corresponds to a formation degree of the Ni2 macrochelate of 37 ± 26%; this contrasts with the one for the other metal ions which amounts on average to 64 ± 9% (3σ) [143]. Upon deprotonation of the phosphate group, which then becomes the primary binding site, the metal ions show their common individual properties. Macrochelate formation according to Equilibrium (24) involving N7 [and possibly also (C6)O)] results for the M(XMP – H) complexes, also written as (X – H • MP • M) [143]. The stability enhancement determined for Ni(XMP – H) is highest among all the M(XMP – H) species studied (for the M2 ions see above). This result is in agreement with the observation made for the M(AMP), M(IMP), and M(GMP) complexes (see Table 5). The stability enhancement, log D Ni(XMP – H) 1.67 ± 0.17 (defined by analogy to Eq. 30), corresponds to a formation degree of 98 ± 1% for (X – H • MP • Ni) cl.

6.4. Flavin Mononucleotide Complexes Flavin mononucleotide (FMN2), also known as riboflavin 5-phosphate, is composed of the 7,8-dimethyl isoalloxazine (i.e., the flavin) ring system, which is bound via N10 to the methylene group of the sugar-related ribit, giving 7,8-dimethyl-10-ribityl-isoalloxazine, also known as riboflavin or vitamin B2 [147], which carries at its 5 position a phosphate monoester residue (see Figure 11, bottom). The corresponding P(O)2 (OH) group has an acidity constant of the expected order, i.e., pKa 6.18 ± 0.01 [148]. Surprisingly, the stability constants of the M(FMN) complexes, where M2 Mg2, Ca2, Sr2, Ba2, Mn2, Co2, Ni2, Cu2, Zn2, and Cd2, are throughout slightly more stable than is expected from the basicity of the phosphate group. Met. Ions Life Sci. 2, 109–180 (2007)


149

This stability enhancement amounts on average to log DM/FMN 0.16 ± 0.04 (3σ) (defined by analogy to Equation 30) including the value for the Ni(FMN) complex, log D Ni/FMN 0.11 ± 0.09 (3σ) [148]. This slight stability increase cannot be attributed to the formation of a seven-membered chelate according to Equilibrium (1) (see Section 2.3) involving the ribit-hydroxyl group at C4. This is demonstrated by the stabilities of the M(G1P) complexes which contain the same structural unit (see Figure 11), but show no enhanced stability, as already discussed in Section 2.3. Hence, in agreement with the results of a kinetic study [149] one has to conclude that the slight stability increase of the M(FMN) complexes has to be attributed to the isoalloxazine ring. However, the equality of the stability increase of the complexes for all ten metal ions mentioned precludes its attributions to an interaction with a N site of the neutral isoalloxazine ring and makes a specific interaction with an O site also rather unlikely (note, deprotonation of (N3)H of FMN2 occurs only with pKa 10.08 ± 0.05 [148]). Furthermore, carbonyl oxygens in aqueous solution are not favored binding sites in macrochelate formation, as was concluded in Section 2.3. Maybe an outersphere interaction could be proposed – but to which site? The most likely explanation is the earlier proposal [148] that the small, but overall significant stability increase originates from ‘bent’ M(FMN) species in which the hydrophobic flavin residue [150] is getting close to the metal ion at the phosphate group, thereby lowering the intrinsic dielectric constant (permittivity) in the microenvironment of the metal ion and thus promoting indirectly the PO32M2 interaction [148]. It may be added that the above-mentioned average stability enhancement, log DM/FMN 0.16 ± 0.04, results in KI 0.45 ± 0.13 (3σ) and a formation degree of the ‘bent’ M(FMN) species of 31 ± 6% (3σ) [148]. It is amazing to observe that this average formation degree is, within the error limits, identical with the value calculated [148] for Ni(FMN) cl, i.e., 38 (±10)%, based on kinetic results provided in [149]. Moreover, the stability data of Stuehr et al. [149] lead to log DNi/FMN 0.17 and yield 32% for Ni(FMN) cl. To conclude, though the given explanation for the enhanced stability of the M(FMN) complexes remains somewhat uncertain, the occurrence of such a stability enhancement for the M(FMN) complexes including Ni2 is definite.

7.

COMPLEXES OF SOME NUCLEOTIDE DERIVATIVES AND ANALOGS

The use of structurally altered nucleotides as probes is one way to study reactions of enzymes which involve nucleotides as substrates. Another goal for structural alterations is the hope of obtaining compounds with useful pharmaceutical properties. In fact, over the years all three parts of nucleotides have been systematically modified, i.e., the nucleobases, the ribose, and the phosphate residues [151,152]. In this section we shall discuss at least one example each by concentrating on such derivatives for which also the Ni2 complexes have been studied. Met. Ions Life Sci. 2, 109–180 (2007)

150

SIGEL and SIGEL

7.1. Complexing Properties of 1,N6 -Ethenoadenosine and of Its Phosphates Formation of a 1,N6-etheno bridge at the adenine residue leads to a 1,10-phenanthroline-like metal ion binding site in adenosine and gives so-called ε-adenosine (ε-Ado) [152]. It is thus no surprise that the stability of the Ni2 complexes increases by a factor of nearly 100 by going from adenosine (log KNi Ni(Ado) 0.4 ± 0.2; Table 1) to ε-adenosine (log K Ni 2.2 [153]). It may be added that N6 of Ni(ε-Ado) ε-Ado can be protonated and that this proton is released with pKH H(ε-Ado) 4.05 ± 0.01 [154]. For the most likely resonance structure of H(ε-Ado) see ref. [154]. In accord with this are the acid–base properties of H2 (ε-AMP) ± (for the structure see Figure 12, top [155,156]) which releases the proton from (N6)H with pKHH2 (ε-AMP) 4.23 ± 0.02 and the one from P(O)2 (OH) with pKHH(ε-AMP) 6.23 ± 0.01 [109,157]. Hence, in the neutral pH range complexes of the type M(ε-AMP) are expected and actually observed [109,157] though M(H; ε-AMP) species are also known [157]. The stability enhancement of the Ni(ε-AMP) complex, compared with a sole Ni2-phosphate binding, amounts to approximately 2 log units, meaning that the macrochelated species (Equilibrium 24) dominate with a formation degree of about 99% [109]. In Section 5.3 we have seen that Ni(ATP)2 experiences a stability enhancement of log D Ni/ATP 0.36 ± 0.06 due to N7 binding of the phosphate-coordinated Ni2 and that the macrochelate reaches a formation degree of about 55% (Table 7). This result needs to be compared with that for Ni(ε-ATP)2 where the stability enhancement amounts to log D Ni/ε-ATP 1.5 and the formation degree 2 of Ni(ε-ATP) 2 cl to 97% [158]. The Ni -promoted hydrolysis of ε-ATP has not been studied, but from the results obtained for the Cu2ε-ATP and Zn2ε-ATP systems [159] it is clear that the dephosphorylation reactions of ε-ATP differ significantly from those of ATP [26]. For example, the reaction in the Cu2ATP system proceeds via a dimeric complex, as the dependence of the reaction rate on the square of the concentration shows [26], whereas in the hydrolysis of the Cu2ε-ATP system only a monomeric species is involved [159]. To conclude, from these results it is evident that great care needs to be exercised if ε-adenosine phosphates are employed as probes for adenosine phosphates in the presence of divalent 3d-transition metal ions.

7.2. Complexes of (N1)-Oxides of Adenosine and Inosine Nucleotides Adenosine forms rather weak complexes with the divalent metal ions of the second half of the 3d-transition series [65,66] (Table 1). However, its (N1)oxidation leads to adenosine (N1)-oxide (Ado • NO) and this nucleoside derivative has strongly enhanced coordinating properties [160]: (N1)-oxidation facilitates Met. Ions Life Sci. 2, 109–180 (2007)


ε -AMP2– 7

5

– –

O

O

O

P O

O H

–

O

N

5

O P

O

O

4 3

9

5'

CH2

N O

6

N

O H

O

–

7

O

–

HO

N O O

P O

2'

9

5'

CH2 4'

H(IMP . NO)–

O H

(AMP. NO)2– O H

1N

2'

1'

4' 3'

+

N

3'

O H

5 4

N O

2–

O S

P

O

O

UMPS2–

4

1' 2'

O H

1

5'

N

CH2 O 3'

O H

2'

O H

7

N

2 9

O

–

1'

4'

NH2

PMEA2–

3

NH

–

O

N

O P O

C H2

O

3

N

O 5

6

H – + O N 1

...

NH2 7

10 1N

1'

4' 3'

–

N O

6

4 3

9

5'

CH2

11

6

N N

151

C H2

5 4

6

N

1 3

N

CH2

Figure 12. Chemical structures of 1,N6-ethenoadenosine 5-monophosphate (ε-AMP2), of adenosine 5-monophosphate (N1)-oxide, (AMP •NO)2, of the monoprotonated form of inosine 5-monophosphate (N1)-oxide, H(IMP • NO) , and of uridine 5-O-thiomonophosphate (UMPS2), as well as of the dianion of the acyclic nucleotide analog 9-[2-(phosphonomethoxy)ethyl]adenine (PMEA2). To facilitate comparisons between ε-AMP2 and AMP2 the conventional atom numbering for adenines is adapted, a procedure which is common [152]. The three purine-nucleotide derivatives are shown in their dominating anti conformation [48,63,107]; the same is true for the pyrimidine-nucleotide derivative UMPS2 [48,108]. 1H-NMR shift measurements have shown [155] that in solution PMEA2 adopts an orientation which is similar to the anti conformation of AMP2; this conclusion is in accord with a crystal structure study [156] of the H2(PMEA) ± zwitterion.

deprotonation of the amino group from pKHAdo 16.7 [161] to pKHAdo • NO 12.86 [160] and the resulting deprotonated o-amino (N1)-oxide group is an excellent chelator [160,162]. Consequently, the nucleotide derivative adenosine 5-monophosphate (N1)-oxide, (AMP • NO)2, offers metal ions two different binding Met. Ions Life Sci. 2, 109–180 (2007)

152

SIGEL and SIGEL

sites, namely the phosphate group (Section 4) and the deprotonated o-amino Noxide unit (Figure 12; middle, left). For steric reasons a simultaneous binding of the same metal ion to both sites is impossible. It may also be added that the neutral o-amino N-oxide group forms only very weak complexes [163]. The acidity constants of H(AMP • NO) are pKHH(AMP • NO) 6.13, corresponding to the deprotonation of the phosphate group, and pKHH(AMP • NO) 12.49, quantifying the deprotonation of the o-amino N-oxide group [164]. The stability constants of the Ni2 complexes are for Ni(AMP • NO), log KNi Ni(AMP • NO) 2.66, and for Ni(AMP • NO – H) , log KNi 7.45 [164]. The first value corresponds Ni(AMP • NO – H) to the stability constant of the Ni(AMP) complex (see Table 5) and the second is close to the stability constant of the Ni(Ado • NO – H) species, log KNi Ni(Ado • NO – H) 7.52 [160]. Of course, application of the mentioned equilibrium constants allows calculation [165] of the acidity constant of the deprotonation reaction of the Ni(AMP • NO) complex: Ni(AMP•NO)

Ni (AMP•NOH)

H K Ni(AMP • NO) [ Ni(AMP•NO H) ][ H ][ Ni(AMP •NO)]

(31a) (31b)

The corresponding acidity constant pKHNi(AMP • NO) 7.70 means that at pH 7.70 the metal ion changes its place from the phosphate group to the deprotonated o-amino N-oxide site. At this pH the affinities of both sites for Ni2 are equal; at pH 7.7 the phosphate group is favored, and at pH 7.7 the o-amino N-oxide group dominates. From the above it follows that the proton and the metal ion compete for binding at a given site. Because the two acidity constants of H(AMP • NO) are so different, the competition of the proton for the two sites is also different. However, this effect can be taken into account by calculating apparent stability constants, Kapp, which are of course no longer universal constants, but are valid only for the pH for which they are calculated [93,164]. The relationship between the stability constant of the complex, ML, the acidity constant of the ligand, HL, and the pH of a solution is given for monobasic binding sites by Equation (32): M K app K ML

1 H 1[H ]K HL

(32)

Application of Equation (32) to the above-mentioned equilibrium constants leads to the situation depicted in the left part of Figure 13. It becomes thus obvious that, e.g., the apparent affinity of the phosphate group for Ni2 is about 1.6 log units larger at low pH values than that of the deprotonated o-amino Noxide group. It should be noted, however, that this alone says little about the absolute concentration of the 1:1 complex formed, but it means that under these conditions Ni2 binds almost exclusively to the phosphate group. Increasing pH favors the metal ion affinity of the o-amino N-oxide group more than that of the Met. Ions Life Sci. 2, 109–180 (2007)


153

Figure 13. Comparison of the metal ion affinity of the phosphate group on the one hand and of the deprotonated o-amino N-oxide unit on the other in dependence on pH for the Ni2 and Cu2 systems. The metal ion affinity is expressed in each case by the apparent stability constants calculated with Equation (32) and the equilibrium constants mentioned in the text (Ni2); for the Cu2 systems the values given in [164] were used. This figure is an altered version of Figure 5 published in [164] (altered by permission of the Verlag Helvetica Chimica Acta, Zürich).

phosphate group. The ‘switch’ of Ni2 from the phosphate to the deprotonated o-amino N-oxide unit occurs at the crossing point of the two curves, i.e., where pH pKHNi(AMP • NO) 7.70; these curves represent the apparent metal ion affinity of the two binding sites in their dependence on pH. The corresponding pKHM(AMP • NO) values of the M(AMP • NO) complexes of Mn2, Co2, and Zn2 are 8.92, 7.77, and 6.90, respectively [164]. For Cu2 the situation is different (Figure 13, right) because in both complexes, Cu(AMP • NO) and Cu(AMP • NO – H) , the metal ion is located at the deprotonated o-amino N-oxide group, which means that in Cu(AMP • NO) the phosphate group carries a proton that is lost with pKHCu(AMP • NO) 5.44 [164]. From the above discussion it is obvious that in many instances it is possible to estimate the coordination tendency of a certain binding site in an ambivalent ligand from the metal ion affinity of the corresponding monovalent ligand offering only this site. With this reasoning in mind, one may consider, e.g., the Ni2 binding properties of adenosine 5-diphosphate (N1)-oxide. The stability of the phosphate residue may then be represented by the value for Ni(ADP) (Table 6) and the one of the deprotonated o-amino N-oxide group by that of Ni(AMP • NO – H) (see above). Application of these equilibrium constants gives for pKHNi(ADP • NO) 8.97 and this means that the ‘switching’ point of Ni2 from the phosphate residue to the deprotonated o-amino N-oxide unit occurs at pH 8.97. Indeed, strengthening Met. Ions Life Sci. 2, 109–180 (2007)

154

SIGEL and SIGEL

of the phosphate–metal ion affinity must shift the ‘switch’ of Ni2 to higher pH values and this is observed. For the monophosphate it is at pH 7.7 ( pKHNi(AMP • NO)) and for the diphosphate at 9.0. Naturally, if the apparent stability constants of the two binding sites become similar then both isomers occur in equilibrium with each other. This is actually the case for the Cu2/ADP • NO3 system in the pH range below 6.2 ( pKHCu(ADP • NO)); thereafter, with increasing pH, Cu2 binding at the deprotonated o-amino N-oxide group will increasingly be favored [165]. The substitution of the amino group in adenosine (N1)-oxide nucleotides by a hydroxy group leads to inosine (N1)-oxide nucleotides [166] and hence, to another set of ambivalent ligands [167]. The structure of the most simple one, i.e., inosine 5-monophosphate (N1)-oxide, IMP • NO2, is shown in Figure 12 (middle; at the right). Possible tautomeric forms of the modified nucleobase residue have been discussed [168], but they are of no relevance here because remarkable metal ion binding occurs only in the deprotonated form. The acidity constants of H(IMP • NO) are pKa 5.43 and 6.31 whereby the first proton is largely released from the o-hydroxy N-oxide unit and the second one from the P(O) 2 (OH) group [168]. The micro stability constants of the Ni(IMP • NO – H) species have been estimated, based on the measured stability constants [167], as being log kNi Ni(NO – H) 4.2 for binding to the deprotonated o-hydroxy N-oxide site and log kNi Ni(PO) 2.8 for binding to the phosphate group [165]. From this follows that at pH 8, where competition with the proton no longer occurs, in the 1:1 complex about 4% of Ni2 are phosphate-coordinated and about 96% are at the nucleobase. As discussed above for AMP • NO, phosphorylation of IMP • NO2 to give IDP • NO3 will favor phosphate-Ni2 binding. If one estimates log kNi Ni(POPO) 4.0 for the diphosphate group (Table 6), one obtains for pH 8 that in the 1:1 complex about 40% of Ni2 are phosphate-coordinated and about 60% are at the deprotonated o-hydroxy N-oxide unit. Application of Equation (32) allows one again to take the competition of protons at the two binding sites into account and to calculate the intramolecular metal ion distribution also at lower pH values [165,167].

7.3. Complexes of Nucleoside 5-O-Thiomonophosphates The phosphate group of nucleotides has been altered in many ways [151]. Probably the most popular alteration is to substitute one of the oxygens by a sulfur atom. Such derivatives are then commonly employed as probes in ribozymes [102,169]. Nevertheless, quantitative information on Ni2 binding to OP(S)(O)22 groups is scarce. There are only two studies, one involving adenosine 5-O-thiomonophosphate (AMPS2) [151] and the other uridine 5-O-thiomonophosphate (UMPS2) (see Figure 12; bottom, left) [170]. Into the latter study methyl thiophosphate (MeOPS2) was also included for reasons of comparison. Thereby it was proven, Met. Ions Life Sci. 2, 109–180 (2007)


155

in accord with the discussion in Section 5.2.2, that the uracil residue is not involved in metal ion binding in M(UMPS) complexes. It may be added that thiophosphates are considerably less basic than phosphates with pKa values of about 4.8 [170,171]. It was further concluded that one of the two negative charge units in a thiophosphate group is located on the sulfur atom whereas monoprotonation occurs at one of the two terminal oxygen atoms [171]. The stability constants of the Ni(PS) complexes, where PS2 MeOPS2, UMPS2 or AMPS2, are listed in column 2 of Table 8, together with the corresponding data for the Mg(PS) and Zn(PS) species. For the M(AMPS) complex it is assumed that macrochelate formation occurs in the same extent as in the M(AMP) complexes (see Section 5.2.3). In this way the stability enhancement due to macrochelation can be corrected for (Table 8, column 3) and one

Table 8. Stability constant comparisons for the M(PS) complexes formed by Mg2, Ni2, or Zn2 and methyl thiophosphate (MeOPS2), uridine 5-O-thiomonophosphate (UMPS2) or adenosine 5-O-thiomonophosphate (AMPS2) between the measured stability constants, log KM M(PS) (Equation 29), or the stability constants of M(AMPS) corM/cor rected for the macrochelate effect, log KM(AMPS) , and the calculated stability constants, M log KM(PS)calcd , together with the stability difference (Equation 33) for aqueous solutions at 25C and I 0.1 M (NaNO3).a,b M(PS)

log KM M(PS)

Mg(MeOPS) 1.33 ± 0.07 Mg(UMPS) 1.24 ± 0.05 Mg(AMPS) 1.28 ± 0.04 Ni(MeOPS) Ni(UMPS) Ni(AMPS)

1.62 ± 0.05 1.54 ± 0.08 2.35 ± 0.07

Zn(MeOPS) Zn(UMPS) Zn(AMPS)

2.34 ± 0.05 2.21 ± 0.06 2.52 ± 0.18

log D M/AMPc

0.06 ± 0.05

0.61 ± 0.06

0.25 ± 0.09

d M/cor log KM(AMPS) log KM M(PS)calcd

log D M/PS

1.22 ± 0.06

1.30 ± 0.03 1.27 ± 0.03 1.25 ± 0.03

0.03 ± 0.08 0.03 ± 0.06 0.03 ± 0.07

1.74 ± 0.09

1.64 ± 0.05 1.59 ± 0.05 1.58 ± 0.05

0.02 ± 0.07 0.05 ± 0.09 0.16 ± 0.10

2.27 ± 0.20

1.69 ± 0.06 1.63 ± 0.06 1.61 ± 0.06

0.65 ± 0.08 0.58 ± 0.08 0.66 ± 0.21

a

For the error limits see footnotes a of Tables 1 and 2. The values for AMPS are from [151], those for MeOPS and UMPS from [170]. b The acidity constants are for H(MeOPS) pKHH(MeOPS) 4.96 ± 0.02 (Eq. 26), for H(UMPS) pKHH(UMPS) 4.78 ± 0.02 (Eq. 26) and pKHUMPS 9.47 ± 0.02 (Eq. 27) [170], and for H2 (AMPS) ± pKHH2 (AMPS) 3.72 ± 0.03 (Eq. 25) and pKHH(AMPS) 4.83 ± 0.02 (Eq. 26) [171]; the micro acidity conAMPS•H stants for H2 (AMPS) ± , the second one being needed for the straight-line calculation,d are pkH•AMP•H AMPS 3.84 ± 0.02 [(N1)H deprotonation] and pkAMPS•H 4.71 ± 0.04 (deprotonation of the thiophosphate group) [171]. c See Table 5 in Section 5.2.3 or [151]. d The values in this column were calculated with the pKHH(PS)values given aboveb by analogy to Equation (4a) by using the straight-line parameters listed in [25,53,55].

Met. Ions Life Sci. 2, 109–180 (2007)

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SIGEL and SIGEL

obtains then stability values which reflect only the influence of the sulfur atom on metal ion binding in M(AMPS) species (column 4). Application of the previously defined straight-line plots (see Figure 9) allows then, together with the acidity constants of the H(PS) species (see footnote b of Table 8), calculation of the stability of the complexes due to a (thio)phosphate-M2 coordination (Table 8, column 5). Together with these values the stability difference according to Equation (33) can be calculated and in this way the effect of the sulfur substitution can be quantified: M M log DM PS log K M(PS) log K M(PS)calcd

(33)

From the results listed in column 6 of Table 8 it is evident that there is no difference in stability between the Mg2 complexes of phosphate (R-PO32) or thiophosphate monoesters (PS2) if the different basicities of R-PO32 and PS2 are taken into account. On the other hand, all three Zn(PS) complexes show, within the error limits, the same enhanced complex stability proving that the S atom of the thiophosphate group participates in Zn2 binding. For the Ni(PS) complexes no unequivocal conclusion can be made. From the data for Ni(MeOPS) and Ni(UMPS) it appears that sulfur substitution has no effect on complex stability whereas for Ni(AMPS) a small effect is indicated; maybe the extent of macrochelate formation in Ni(AMP) and Ni(AMPS) is slightly different. In any case, by applying the procedure described in [151] and [170] one calculates for the Zn(PS) complexes from the average value, log D Zn/PS 0.63, that about 77% of Zn2 is S-coordinated and 23% is O-coordinated to the –OP(S)(O)22 residue. Assuming for Ni(PS) an upper limit of 0.1 for log D Ni/PS, up to about 20% of the complexes may be S-coordinated. Here clearly more research is needed. However, overall it is clear that Zn2 is more thiophilic than Ni2 and this is in accord with the solubility products of ZnS and NiS [151].

7.4.

Complexes of Acyclic Nucleotide Analogs

One way to alter the cyclic ribose residue of a nucleotide is to ‘delete’ it. Indeed, acyclic nucleotide phosphonates, i.e., analogs of (2-deoxy)nucleoside 5-monophosphates, have been increasingly studied during the past two decades. One of these compounds, 9-[2-(phosphonomethoxy)ethyl]adenine (PMEA; Adefovir; see Figure 12, bottom, right) was approved in 2002/2003 in the form of its bis(pivaloyloxymethyl)ester for use in the therapy of hepatitis B, a disease evoked by a DNA virus. Diphosphorylated PMEA2, i.e., PMEApp 4, is initially recognized by nucleic acid polymerases as an excellent substrate, but after insertion into the growing nucleic acid chain, transcription is terminated due to the lack of a 3-hydroxyl group. Based on the metal-ion-binding properties of PMEApp 4 it can be explained why the ether oxygen in the aliphatic chain, RCH2OCH2PO3pp 4, is compulsory Met. Ions Life Sci. 2, 109–180 (2007)


157

for a useful biological activity. The systematic progress made over the past 15 years [53] in understanding the coordination chemistry of these compounds can be followed in the reviews [172] (1995), [173] (1999), and [174] (2004). It can be seen from Figure 9 that the metal-ion-binding properties of PMEA2 and AMP2 differ, if one considers the stability of the Mg2 complexes. Mg(PMEA) is evidently more stable than Mg(AMP)! By inclusion of the dianion of (phosphonylmethoxy)ethane (PME2) it could be shown that this increased stability is solely due to the formation of a five-membered chelate involving the ether oxygen as expressed in Equilibrium (34):

R

O

H C H

H2 C

O P O– O–

R

O PO– O–

O

M2+

(34)

M2+

The percentage of the closed species amounts for Mg(PMEA) to 31 ± 8% [53]. For Mn(PMEA) and Zn(PMEA) the situation is quite similar and the enhanced complex stabilities, log DMn/PMEA 0.21 ± 0.08 and log D Zn/PMEA 0.30 ± 0.10 (Eq. 30), are solely explained with Equilibrium (34), the corresponding formation degrees of the closed species being 38 ± 11 and 50 ± 12% [53,173]. However, detailed studies over the years revealed that for Cu2 [175] and Ni2 [176] the situation is much more complicated and that the four M(PMEA) isomers indicated in Scheme (35) are in equilibrium with each other:

M(PMEA)cl/N7

2+

M

+ PMEA

2–

M M(PMEA)op

I/N7

M(PMEA)op I/O

(35)

M(PMEA)cl/O I/O/N3

M(PMEA)cl/O/N3

This means, there is an ‘open’ species in which M2 is only phosphonatecoordinated, M(PMEA) op, and which then interacts either with N7 or the ether O atom giving the chelated isomers, M(PMEA) cl/N7 or M(PMEA) cl/O, respectively. Finally, the latter species may, in addition to the five-membered chelate, also form a seven-membered ring by binding the coordinated M2 also to N3, Met. Ions Life Sci. 2, 109–180 (2007)

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M(PMEA) cl/O/N3. The formation degrees of the four isomers M(PMEA) op, M(PMEA) cl/N7, M(PMEA) cl/O, and M(PMEA) cl/O/N3 are about 17, 8, 34, and 41%, respectively, for the Cu2 system [175–177] and about 50, 13, 19, and 18%, respectively, for the Ni2 system [176]. Evidently, the dominating isomers in the case of Cu2 are those involving the ether O. In fact, the two isomers, Cu(PMEA) cl/O and Cu(PMEA) cl/O/N3 amount together to 75%. For the Ni2 system the open isomer dominates with about 50%, but the two isomers involving the ether O amount together still to about 37%. These results confirm the rather high stability of the five-membered chelates already indicated above in Equilibrium (34). In contrast, the corresponding sixmembered chelate that could be formed with the dianion of 9-[2-(phosphonoethoxy)ethyl]adenine (PEEA2) [one more CH2 group is inserted between O and PO32] is considerably less stable. With Mg2 the chelate does not form at all, log DMg/PEEA 0.01 ± 0.07 [178], and in the Ni2 system M(PEEA) cl/O is with about 4% a minority species, possibly not occurring at all. Instead M(PEEA) cl/N7 reaches a formation degree of about 38%. But the dominating species is clearly the open isomer Ni(PEEA) op with 58%. Similarly, with Cu2 the stability of the six-membered chelates is also smaller than that of the five-membered ones [178]. Two lessons are to be learned here: (i) Five-membered chelates are more stable than six-membered ones, and this observation offers an explanation why PMEA is a nucleotide analog with excellent antiviral properties and PEEA is not. In the latter case the second metal ion needed for the nucleotidyl transfer is not properly adjusted at the triphosphate residue [174,178] (see also the second to the last paragraph in Section 5.3). (ii) If properly pre-orientated by other primary binding sites, N3 of a purine is well able to coordinate metal ions including Ni2 (see also Section 6.1).

7.5. Nickel(II) Binding to Nucleotides Containing a Platinum(II)-Coordinated Nucleobase Residue A further way to alter nucleotides via their nucleobases (cf. Sections 7.1 and 7.2) is to ‘fix’ a kinetically inert metal ion to the nucleobase. Several such examples have been studied on the one hand with cis-(NH3)2Pt2 coordinated to N3 of dCMP2 [115] or to N7 of dGMP2 [179]. The main outcome of these studies is that the acid–base properties of the phosphate groups are only relatively little affected [180] and that this is also true for binding of divalent metal ions to the phosphate residues [115,179]. To give a specific example: Coordination of cis-(NH3)2Pt(dGuo-N7)2 to the N7 site of dGMP2 results in the complex cis-(NH3)2Pt(dGuo-N7)(dGMP-N7) which may be protonated at its phosphate group. Comparison of the corresponding acidity constant, pKHH[Pt(dGuo)(dGMP)] 5.85 ± 0.04, with pKHH(dGMP) 6.29 ± 0.01 reveals an acidification of the P(O)2 (OH) group due to charge repulsion by the N7-coordinated Pt(II) of ∆pKa 0.44 ± 0.04 [181]. The average acidification Met. Ions Life Sci. 2, 109–180 (2007)


159

of cis-(NH3)2Pt2 in the same complex on the two (N1)H sites is somewhat more pronounced (as one would expect) and amounts to ∆pKa/av 0.78 ± 0.11 [181]. However, coordination of Mg2, Cu2 or Zn2 to the phosphate group of cis(NH3)2Pt(dGuo-N7)(dGMP-N7) is inhibited on average only by about 0.2 log unit [181]. The same may be surmised for Ni2 binding. This moderate effect of 0.2 log unit suggests with regard to DNA or RNA that cis-(NH3)2Pt2, if coordinated to N7 of a guanine residue, affects metal ion binding to the nucleic acid phosphate backbone only little. Another interesting case concerns the acyclic nucleotide analogue PMEA2 (Figure 12; bottom left), already discussed in Section 7.4. Coordination of (Dien)Pt2 either to N1 [182] or to N7 [183] leads, if monoprotonated, to the complexes H[(Dien)Pt(PMEA-N1)] and H[(Dien)Pt(PMEA-N7)]. Acidification of the P(O)2 (OH) group by Pt2 at N1 gives ∆pKa/N1 0.21 ± 0.03 (based on pKHH(PMEA) 6.90 ± 0.01) and by Pt2 at N7 ∆pKa/N7 0.44 ± 0.01 [182]. The higher acidity of H[(Dien)Pt(PMEA-N7) provides evidence, that in the (Dien)Pt(PMEA-N7) complex in aqueous solution an intramolecular, outersphere macrochelate is formed through hydrogen bonds between the PO32 residue of PMEA2 and a Pt(II)-coordinated (Dien)-NH2 group. This formation amounts to about 40% and is in accord with several other related observations [182]. The macrochelate mentioned also offers at least a partial explanation for the somewhat different metal ion affinities, including Ni2, of the phosphonate groups of (Dien)Pt(PMEA-N1) and (Dien)Pt(PMEA-N7) which amount on average to log D N1/av 0.17 ± 0.06 and log D N7/av 0.42 ± 0.04 [182]. This inhibition is due to charge repulsion of the adenine-coordinated (Dien)Pt2 and the M2 coordinated at the PO32 group. It needs to be mentioned that, in all these cases, including the Ni2 systems, no indication for the existence of Equilibrium (34) (see Section 7.4) was observed, i.e., the ether oxygen of the ‘aliphatic’ side chain of PMEA2 does no longer participate in M2 binding. The only exceptions here are the corresponding Cu2 and Zn2 complexes where the five-membered chelates still form to some extent [182]. Overall, these results confirm the above conclusion that a purine-coordinated Pt2 unit affects metal ion binding at the phosphate group of the same nucleotide only relatively little. This is of relevance for multiple metal ion binding to nucleotides and nucleic acids (see also the final paragraph of Section 3.1).

8. 8.1.

MIXED LIGAND COMPLEXES CONTAINING A NUCLEOTIDE Some General Comments and Definitions

A mixed ligand or ternary complex of the kind to be considered here is composed of a metal ion and two different coordinated ligands. There are various ways Met. Ions Life Sci. 2, 109–180 (2007)

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to quantify the stability of such ternary complexes [184,185]. First we consider complexes in which a nucleotide ( NP nucleoside phosphate; charges omitted) is bound in the coordination sphere of M2 and to which a second ligand L (charges deleted) is binding, leading thus to the ternary M(NP)(L)2 complex and Equilibrium (36a): M(NP)2 L

M(NP)(L)2

M(NP) K M(NP)(L) [ M(NP)(L)2 ]([ M(NP)2 ][ L])

(36a) (36b)

Evidently, this equilibrium is best compared with the following one: M2 L

M(L)2

(37a)

[ M(L)2 ]([ M2 + ][ L])

(37b)

M(NP) M ∆ log K M NP L log K M(NP)(L) log K M(L)

(38a)

M K M(L)

The difference ∆ log KM/NP/L,

M(L) M log K M(L)(NP) log K M(NP)

(38b)

characterizes the coordination tendency of L towards M(NP)2 (Eq. 36) relative to M(aq)2 (Eq. 37) and vice versa (Eq. 38b) [184,185]. The latter point is important because it means that the difference ∆ log KM/NP/L (Eq. 38) also equals the one resulting from the following two equilibria: M(L)2 NP

M(NP)(L)2

M(L) 2 2 K M(NP) ( L) [ M(NP)(L) ]([ M(L) ][ NP])

M2 NP M K M(NP)

M(NP)2 [ M(NP)2 ]([ M2 ][ NP])

(39a) (39b) (40a) (40b)

Factors which arise through direct [114,186,187] or indirect (i.e., metal-ionmediated) [57,184,185,188] ligand–ligand interactions, they may be either favorable or unfavorable, will show up in this description. It is important to note that ∆ log KM/NP/L is the difference between two log stability constants and thus has to be a constant itself. Indeed, it quantifies the Met. Ions Life Sci. 2, 109–180 (2007)


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position of Equilibrium (41a): M(NP)2 M(L)2 10 ∆ log K M NP L

M(NP)(L)2 M2 [ M(NP)(L)2 ][ M2 ] [ M(NP)2 ][ M(L)2 ]

(41a) (41b)

Since more coordination positions are available for binding of L to M2 (Eq. 37) than to M(NP)2 (Eq. 36), one expects on statistical grounds [184,185] and in accord M with the general rule [93] that KM(L) KM(L) M(L) 2 negative values for ∆ log KM/NP/L. For example, for the coordination of a bidentate ligand followed by a monodentate one in a regular octahedral (oh) coordination sphere like that of Ni2, statistical considerations [189] provide the value ∆ log K2:1/stat/oh log (4:1/6:1) 0.18. For two different bidentate ligands ∆ log K2:2/stat/oh log (5/12) 0.38 [184] results. Correspondingly, for the Jahn–Teller-distorted octahedral coordination sphere of Cu2 a statistical value is more difficult to assess, but on the basis of previously advanced arguments [184] one obtains the values ∆ log K2:1/stat/Cu 0.5 and ∆ log K2:2/stat/Cu –0.9 [184,189]. The available information on mixed ligand complexes containing Ni2 and a nucleotide is rather limited. However, we shall concentrate below mainly on ternary complexes containing NTPs because in these instances the available equilibrium constants are relatively well defined.

8.2.

Ternary Nickel(II) Complexes Containing ATP 4 and a Buffer Molecule

Considering that many experiments in biochemistry are carried out in buffers to stabilize the pH of a solution, it is important to indicate possible drawbacks of this procedure. In fact, the Zn2-promoted dephosphorylation of ATP is inhibited by acetate, Tris or borate buffers [26,130]. Similarly, buffers composed of imidazole [190] or phosphate [191,192] also inhibit the reactivity of M2/ATP systems. Corresponding effects must be anticipated for Ni2/NTP systems. Tris is one of the most popular buffers used in biochemical studies, probably because its buffer region encompasses the physiological to slightly alkaline pH range [193]. The same is true for Bistris which buffers in the pH range 6 to 7.5 [194]. Another widely employed buffer is Bicine [195,196] which is employed in the pH range 7.5–9. The structures of the three mentioned buffers are shown in Figure 14. Because Bicine is derived from glycine, it was expected already 40 years ago that this buffer forms complexes with metal ions [197]. For Tris and Bistris the awareness that in the presence of metal ions interactions between the two buffers and these ions need to be considered is much lower, and the fact that also mixed ligand complexes may form [194,195], has hardly been realized. Therefore, the Met. Ions Life Sci. 2, 109–180 (2007)

162

SIGEL and SIGEL HO HO

CH2

C

HO

CH2

HO

CH2

CH2

HO

CH2

HO

C CH2

O C

CH2

–O

NH2

Tris

CH2

CH2

OH

Bistris

N CH2

CH2

OH

CH2

CH2

OH

NH

Bicine

+

CH2

CH2

OH

Figure 14. Chemical structures of 2-amino-2-(hydroxymethyl)-1,3-propanediol (Tris), 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)-1,3-propanediol (Bistris), and N,Nbis(2-hydroxyethyl)glycine (Bicine).

stabilities of the ternary Ni2 and some related other M2 complexes formed between these buffers and ATP 4 are briefly summarized. The stability constants according to Equilibria (36a) and (37a) are listed in Table 9 together with the stability differences defined in Equation (38). The stabilities of the binary M2 complexes of Tris and Bistris are quite large (Table 9, column 4) and this fact has been attributed to chelate formation with the hydroxyl groups [193,194]. The significant role of the hydroxyl groups of these buffers is also confirmed by a comparison of the stability constants of the M(Tris)2 complexes with those of the corresponding M(Bistris)2 species (Table 9, column 4). The stability increase is between 0.05 and 1.2 log units despite the fact that the basicity of the nitrogens in Bistris is about 1.5 pK units lower (see footnote b in Table 9). This proves the earlier conclusion that most of the hydroxyl groups of Bistris participate in metal ion binding [194]. The hydroxyl groups also play an important role in the ternary complexes as is evident from the small negative values observed for ∆ log KM/ATP/L (Table 9, column 5). Indeed, the role of the hydroxyl groups for the stability of the M(ATP)(Tris)2 complexes follows from comparisons with the stability conNi(ATP) stants of the M(ATP)(NH3)2 complexes for Ni2 (log KNi(ATP)(NH

2.3 [198]) 3) Cu(ATP) and Cu2 (log KCu(ATP)(NH3) 3.4 [198]). These constants are somewhat smaller than those of the corresponding M(ATP)(Tris)2 complexes (Table 9, column 3) despite the much lower basicity of Tris (pKHH(Tris) 8.13 [193]) compared with that of NH3 (pKHNH4 9.38 [198]). It follows clearly that the OH groups play a role, but it is unclear to which extent they participate in ternary complex formation via direct metal ion binding or via hydrogen bonding to the phosphate oxygens of the coordinated ATP 4. Met. Ions Life Sci. 2, 109–180 (2007)


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Table 9. Stability constants of some ternary M(ATP)(L) (Eq. 36) and binary M(L) complexes (Eq. 37), where L Tris [193], Bistris [194], or Bicinate [195], as determined by potentiometric pH titrations [193,194] or spectrophotometric measurements [195] in aqueous solutions at 25C, together with the stability differences according to Equation (38).a,b M(ATP) log KM(ATP)(L)

M log KM(L)

∆ log KM/ATP/L

L

M2

Tris

Co2 Ni2 Cu2 Zn2

1.57 ± 0.05 2.35 ± 0.05 3.50 ± 0.05 1.8 c

1.73 ± 0.02 2.74 ± 0.02 4.05 ± 0.02 1.94 ± 0.03

0.16 ± 0.05 0.39 ± 0.05 0.55 ± 0.05 0.14

Bistris

Co2 Ni2 Cu2 Zn2

1.33 ± 0.03 2.77 ± 0.04 3.62 ± 0.03 2.0 c

1.78 ± 0.03 3.59 ± 0.02 5.27 ± 0.01 2.38 ± 0.03

0.45 ± 0.04 0.82 ± 0.04 1.65 ± 0.03 0.38

Bicinate

Co2 Ni2 Cu2

4.53 ± 0.22 5.44 ± 0.19 6.57 ± 0.32

5.08 ± 0.13 6.02 ± 0.09 8.24 ± 0.09

0.55 ± 0.26 0.58 ± 0.21 1.67 ± 0.33

a

For the error limits see footnotes a of Tables 1 and 2. H The acidity constants and employed ionic strengths (I) are: (i) pKH(Tris) 8.13 ± 0.01; I H 0.1 M, KNO3 [193]. (ii) pKH(Bistris) 6.72 ± 0.01; I 1.0 M, KNO3 [194]. For the acidity conH 6.56 ± 0.04 [194]. stant of H(Bistris) at 25C and I 0.1 M (KNO3) it holds pKH(Bistris) (iii) The acidity constants of H2(Bicinate) as determined by potentiometric pH titration H are pKHH2 (Bicinate) 2.13 ± 0.06 and pKH(Bicinate) 8.33 ± 0.03; I 1.0 M, KNO3 [195]. c This value is an upper limit of the stability constant, but the actual value is expected to be close to this limit [193,194]. b

However, the values for ∆ log KM/ATP/Bistris as observed for the Co2 and Ni2 complexes are relatively small (i.e., not strongly negative) and the coordination spheres of these metal ions are already quite ‘saturated’ by ATP 4 and the nitrogen of Bistris. Consequently, the most plausible explanation for the relatively high complex stability that remains is the additional formation of hydrogen bonds between some OH groups of the coordinated Bistris and phosphate oxygens of the also coordinated ATP 4. Furthermore, to put the observed negative values for ∆ log KM/ATP/L into perspective, it may be helpful to recall that the expected statistical value for a regular octahedral coordination sphere of M2 (Ni2) and the coordination of two different, but simple and symmetrical tridentate ligands amounts already to ∆ log K3:3/stat/oh 1.03 [194]. The stability constants of the binary M(Bicinate) and ternary M(ATP)(Bicinate)3 complexes speak for themselves. These values are so large (Table 9) that Bicine, if used as buffer in the presence of metal ions, will certainly complex a very significant amount of metal ions present. It is further clear that the hydroxyl groups of Bicinate participate in metal ion binding. Met. Ions Life Sci. 2, 109–180 (2007)

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8.3. Mixed Ligand Complexes Containing a Nucleotide and a Further Monodentate or Bidentate Ligand: Release of N7 and Formation of Stacks From the results discussed in Section 5.3 it is evident that the ‘weak’ point in macrochelate formation of M(ATP)2 complexes is the coordination of N7 of the adenine residue. Indeed, by 1H-NMR shift experiments it has been shown for Zn2 and Cd2 that already the formation of the ternary M(ATP)(OH)3 complexes releases N7 [199]. The same may be surmised for Ni(ATP)(OH)3 which forms with pKHNi(ATP)(H2O) 9.4 [12]. Similar observations have been made for Cd(ATP)(NH3)2 [200]. For the ternary complexes of M(ATP)(imidazole)2, where M2 Mn2, Co2, Zn2 or Cd2, it is shown, based on a careful analysis of stability data, that the adenine moiety is released from the coordination sphere [200]. For Ni(ATP)2 and Cu(ATP)2 the analysis shows that addition of imidazole at least reduces, and possibly completely eliminates, the extent of N7 backbinding [200]. However, at the same time there is also evidence that in these M(ATP)(imidazole)2 complexes intramolecular stacking between the purine moiety and the imidazole ring occurs to some extent, i.e., the formation degree of the stacks being of the order of 15–50% [201]. The values of ∆ log KNi/NTP/Bpy (Equation 38) as measured for the ternary systems composed of Ni2, 2,2-bipyridine, and ATP 4 or ITP 4 are relatively high [202], indicating an increased complex stability. Indeed, by spectrophotometry (charge-transfer bands) of the Co2, Ni2, Cu2, and Zn2 systems as well as by 1H-NMR measurements for the Zn2 one, intramolecular stack formation between the aromatic rings of Bpy and the purine residue of ATP 4 or ITP 4 could be proven [203]. In all ternary M(Bpy or Phen)(ATP)2 complexes the formation degree of the intramolecular stack is very large, usually of the order of 90% [114]. Of course, the extent of intramolecular stack formation depends strongly on the steric opportunities provided by the ligands: There are no data for Ni2 complexes but in Cu(Phen)(3AMP/5AMP or 2AMP) the extent of stacking increases within the given series of AMPs from 56 ± 9, via 90 ± 2 to 97.2 ± 0.5% [204] and is also very large in Cu(Phen)(PMEA) and related complexes (about 90%) [187]. In corresponding mixed ligand systems where Bpy or Phen are replaced by amino acids such as tryptophanate or leucinate also intramolecular stacks involving the indole moiety or hydrophobic adducts with the isopropyl residue form [205]. For example, for Ni(AMP)(tryptophanate) , based on stability constant data [206], a formation degree of 57 ± 24% (3σ) was calculated [114] for the intramolecular stack. Interestingly, for the Ni(ATP)(histidinate)3 complex intramolecular stack formation between the purine moiety and the imidazole ring is also anticipated [207]; no indication for such an interaction was found in Ni(dGMP)(histidinate) , though it was concluded that N7 is released from the coordination sphere of Ni2 [208]. Met. Ions Life Sci. 2, 109–180 (2007)


165

It may be noted that aromatic ring stacking can occur in rather complicated structures, e.g., in the sexternary complex formed by the quaternary cis-(NH3)2Pt(2deoxyguanosine-N7)(dGMP-N7) complex and the binary Cu(Phen)2 species the intramolecular stack reaches a formation degree of 50 ± 14% (3σ) [209]. This example demonstrates nicely how the links by metal ions promote intramolecular interactions – and there is no doubt that Ni2 may take over the role played here by Cu2. Finally, this discussion should not distract from the fact that certain ligand combinations, as discovered 40 years ago [210], can give rise to enhanced complex stabilities, i.e., the combination of a heteroaromatic N base and an O donor ligand in combination with a transition metal ion leads to indirect effects being mediated by the metal ion [57,185,188]. In this context the observation that adenosine forms (mainly) through the coordination of N7 a complex with Ni(hydrogen triphosphate)2 is most interesting: Comparison of log KNi(HTP) Ni(HTP)(Ado) 1.36 ± 0.04 [125] with log KNi 0.32 [125] or 0.4 ± 0.2 (Table 1) reveals that Ado prefers Ni(Ado) coordination at Ni(HTP)2, compared to Ni(aq)2, by a factor of about 10. Clearly, the direct and indirect ligand–ligand interactions that occur in mixed ligand complexes and which were described in this section, are of relevance regarding the discrimination and selectivity observed in nature.

9.

NICKEL(II) BINDING IN NUCLEIC ACIDS

In Section 3.1 we have seen that among the neutral nucleobase residues shown in Figure 3 the N7 sites of purines are favored. Among these Ni2 forms the most stable complexes with N7 of the guanine residue (Table 1), commonly also involving a hydrogen bond to (C6)O of a Ni2-bound water molecule (Figure 1). Indeed, the stability constant of Ni(dGuo)2, log KNi Ni(dGuo) 1.53 ± 0.09 (Table 1), is larger than the binding affinity of Ni2 towards monoprotonated D-ribose 5monophosphate, log KNi Ni(H;RibMP) 0.7 [13], and in Section 4 it was concluded that the interaction of Ni2 with phosphate diester groups occurs predominately in an outersphere manner. With the above information at hand it is no surprise that all known crystal structure studies [211–215] of Ni2 complexes formed with DNA-type oligonucleotides (hexa- [214], hepta- [215], and decanucleotides [211–213]) show the indicated N7 coordination to guanines whereby Ni2 selects between different guanines via their steric accessibility, i.e., it prefers exposed guanines [211–215]. These results are consistent with the proposal that the left-handed Z-DNA conformation is stabilized by metal ions, partly because they interact only with guanine-N7 positions that are more exposed than in B-DNA [214]. Therefore, in an appropriate sequence Co2, Ni2, and Zn2 would favor the Z-DNA conformation [214]. Indeed, that Ni2 promotes in aqueous solution the transition of B- to Z-DNA is known [216] and for poly d(GC) it was shown that different anions affect this transition differently; NaCl disfavors this transition (by a factor of 10) Met. Ions Life Sci. 2, 109–180 (2007)

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much more strongly than NaNO3 [216]. This observation cannot be explained by the binding constant of Ni2 to Cl [216]. In the mentioned crystal structures [211–215] it is overwhelmingly Ni(H2O)52 which coordinates innersphere to the guanine-N7 sites and outersphere to (C6)O [211–214] (cf. also Figure 1 in Section 2.2). However, also some minority interactions occur: A Ni2 ion may innersphere bridge two guanines [211,213] (Co2 can do the same [217]), i.e., it may crosslink different duplexes through extrahelical guanines [212]. (ii) Ni2 may also bridge phosphate groups of two neighboring duplexes [213] and such a double Ni2-phosphate interaction may occur via hydrated Ni2 ions [215]. (iii) Guanine(N7)-Ni2-phosphate interactions between two duplexes were also observed with N7 innersphere and the phosphate group outersphere coordinated [215] but that both interactions occur innersphere has also been shown [212]. (i)

The decanucleotide d(CGTATATACG) duplexes are stacked as infinite columns. Figure 15 shows the section which contains the Ni2 ion coordinated to N7 of

Figure 15. Coordination sphere of the Ni2 ion bridging two duplexes formed by the DNA-decanucleotide d(CGTATATACG) [212]. The octahedral Ni2 ion is innersphere coordinated to N7 of a 3-terminal G10, a phosphate oxygen of T5, and four water molecules, one of which forms a hydrogen bond to (C6)O of G10 (not shown). Further water molecules found in the structure are omitted for clarity. Hydrogen bonds involved in base pairing are indicated by dotted lines, and the DNA backbone as grey ribbons. The structure was prepared using MOLMOL [235] and the coordinates from [212], PDB accession code 473D. Met. Ions Life Sci. 2, 109–180 (2007)


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G20, the corresponding (C6)O being outersphere bound. The same Ni2 also forms an innersphere phosphate bridge to T5 of the neighboring duplex [212]. In accord with the above-mentioned solid-state studies, Raman spectroscopy of calf thymus DNA confirmed for aqueous Ni2 systems that Ni2 coordinates mainly to the N7 sites of guanines and also to adenines, but that in addition also outersphere interactions take place with the phosphate groups [218]. Interestingly, stability constant studies in aqueous solution of the DNA model d(pGpG)3 in the presence of Zn2 or Cd2 (and for Ni2 the same may be surmised) show next to M2–PO32 binding also strong interactions with N(7) [219]. It is further of relevance to note here that a crystal structure study [217] of a hexanucleotide shows innersphere Co2 coordination to N7 of a guanine and outersphere binding to the phosphate bridge of the same nucleotide unit. Considering that Ni2 may bridge DNA duplexes, it is no surprise that stable protein-Ni2–DNA complexes have been found [220] and that the Ni2-mediated assembly of an RNA-amino acid complex was observed [221]. Furthermore, that the ring of a netropsin drug may stack with an unpaired guanine residue in the minor groove of a duplex formed by decanucleotides [212] is also no surprise in the light of the stacking interactions discussed in Section 8.3. Moreover, Ni2 immobilized on a Nta resin (i.e., nitrilotriacetate fixed to a resin with coordinated Ni2) may be used for the in vitro selection of Ni2-binding RNA molecules [222]. By 1 H-NMR spectroscopy and using Ni2 as a paramagnetic probe, it was concluded that the binding site in the selected RNA is near a G-A base pair [222]. In attempts to prepare site-specific agents for the cleavage of DNA or RNA Ni2 complexes have received much attention. For example, it was shown that square planar macrocyclic Ni2 complexes, after oxidation to Ni3 with an octahedral coordination sphere, interacted preferably with N7 of extrahelical guanine residues. This means that addition of oxidizing agents like KHSO5 leads to Ni3 which resulted in guanine binding and oxidation and finally (after piperidine treatment and heating) in strand scission [223]. Similar observations were made with Ni2-salen complexes, the salens being formed from two salicylaldehydes and a diamine [224]. This has further led to the template-directed assembly of metallosalen-DNA conjugates also containing (at least initially) Ni2 [225]. Another closely related approach uses Ni2 tripeptides, like Gly-Gly-His or ArgGly-His, because this also leads (after deprotonation of two peptide protons) to square-planar Ni2 complexes in which the Ni2 may then be oxidized to Ni3; this procedure allows then selective recognition and cleavage of RNA and DNA [224,226]. Much effort has recently been spent on the design of microelectronic circuits to be used in the nanotechnology of the future. These efforts have led to the construction of so-called M-DNA. This is a complex between Co2, Ni2 or Zn2 and duplex DNA which forms at a pH of about 8.5. The preferred model is that in this complex the imino protons of T and G, i.e., of the (N3)H and (N1)H sites, respectively, have been replaced by the metal ions. There are indications that Met. Ions Life Sci. 2, 109–180 (2007)

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M-DNA might to some extent behave as a molecular wire [227] and there are also speculations that under certain conditions M-DNA might have a biological role [228]. In accord with the above is the fact that the formation of M-DNA is strongly pH-dependent [229]. On the other hand, in a study using atomic force microscopy no evidence for metallic or semiconductor behavior was found [230]. It is evident that much more work is needed to delineate the properties of M-DNA and to see if it has properties useful in nanotechnology. Of interest in this context is also the recent synthesis of 2-deoxyribosyl-N1[6-(2-pyridyl)-pyrimidinone ( 4-(2-pyridyl)-pyrimidinone deoxyriboside) [231] and of 2-deoxyribosyl-N9-[6-(2-pyridyl)-purine] [232]. The first of these compounds is formally derived from the natural nucleobase cytosine by replacement of its 4-amino group with pyridine. Similarly, in the second one the (C6)NH2 group of adenine is also replaced with a pyridyl group. The result is in both instances a 2,2-bipyridine-type structure which is an excellent site for Ni2 binding. Indeed, incorporation of each of these derivatives into complementary dodecamers leads to self-pairing in the presence of Ni2 and to stable DNA duplexes [231,232].

10. CONCLUDING REMARKS From the material summarized in this chapter it is evident that the interaction of Ni2 with the three constituents of nucleotides and nucleic acids, i.e., the phosphate, sugar, and nucleobase residues, is a delicate and rather complicated matter. Very often are weak interactions those which determine the actual structure of a species in solution. The macrochelates formed in Ni2-nucleotide complexes are typical examples of these qualities (Equilibrium 24). In this context it should be noted that isomeric equilibria are easily shifted from one side to the other (Sections 5 and 8.3). For example, the conversion of an ‘open’ form into a chelated species (Eq. 24) with a formation degree of about 20% is connected with a stability difference (or enhancement) of only log D 0.1 (Eqs 30 and 38). In other words, ∆G changes by 0.6 kJ/mol only [52,55]. Such a change in complex stability is easily achieved by a small alteration in the intrinsic dielectric constant (ε ; permittivity); for example, phosphate–metal ion interactions will be stabilized by a decrease in the polarity of the solvent [30]. All this can give rise to discrimination and selectivity. A matter of concern are the outersphere interactions which evidently play an important role in the coordination chemistry of Ni2 as we have seen throughout this chapter. The problem is that these interactions are very often only tentatively described or follow by analogy to crystal structure studies. Their formations in aqueous solutions are up to now only poorly defined and aside from the Ni(ATP)2 system (Section 5.3) hardly any quantitative data are available. Here more work needs to be done. Another difficulty refers to the binding of Ni2 to the triphosphate chain of a NTP 4 [68,107,157]: Is Ni2 always α,β,γ -coordinated Met. Ions Life Sci. 2, 109–180 (2007)


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or does the interaction with the α-phosphate group vary in dependence on macrochelate formation with N7 of a purine residue? In other words, does N7-backbinding initiate a change from an innersphere to an outersphere coordination of Ni2 to the α-phosphate group? In fact, there are more questions: Does the described cross-linking of two DNA duplexes by Ni2 (Section 9) have a biological role? Are there any beneficial Ni2nucleotide and/or Ni2-nucleic acid interactions? That nickel-DNA interactions have a role in carcinogenesis is long known [8,233,234] and is discussed in detail in the two terminating chapters of this book.

ACKNOWLEDGMENTS Financial support from the Swiss National Science Foundation (SNF-Förderungsprofessur to R.K.O.S., PP002-68733/1), the Universities of Zürich (R.K.O.S.) and Basel (H.S.), and within the COST D20 programme from the Swiss State Secretariat for Education and Research (H.S.) is gratefully acknowledged, as is the help of Astrid Sigel in the preparation of this manuscript and of Dr Eva Freisinger with regard to Figure 15.

ABBREVIATIONS AND DEFINITIONS The abbreviations for the nucleobases and the nucleosides are defined in Figure 3 and the labeling systems for the di- and triphosphate residues in Figures 6 and 8. Other abbreviations are defined below. AcP2 ADP3 ADP • NO3 AMP2 AMP • NO2 AMPS2 ATP 4 Bicine Bistris Bpy CDP3 3,5-cGMP2 CMP2 CTP 4 cyclam

acetyl phosphate (Figure 2) adenosine 5-diphosphate adenosine 5-diphosphate (N1)-oxide (analogous to Figure 12) adenosine 5-monophosphate adenosine 5-monophosphate (N1)-oxide (Figure 12) adenosine 5-O-thiomonophosphate adenosine 5-triphosphate (Figure 8) N,N-bis(2-hydroxyethyl)glycine (Figure 14) 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)1,3-propanediol (Figure 14) 2,2-bipyridine cytidine 5-diphosphate guanosine 3,5-cyclic monophosphate cytidine 5-monophosphate cytidine 5-triphosphate (Figure 8) 1,4,8,11-tetraazacyclotetradecane Met. Ions Life Sci. 2, 109–180 (2007)

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dCMP2 dGMP2 d(GpG) dGuo DHAP2 Dien d(pGpG)3 dTDP3 dTMP2 dTTP 4 f

ε-Ado ε-AMP2

En 9EtG FMN2 G1P2 GDP3 GMP2 GTP 4 HTP3 I IDP3 IDP • NO3 IMP2 IMP • NO2 ITP 4 Ka L M2 MBI 9MeA 9MeG 9MeHyp MeOPS2 MIm NB NDP3

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2-deoxycytidine 5-monophosphate 2-deoxyguanosine 5-monophosphate 2-deoxyguanylyl(3→5)-2-deoxyguanosine 2-deoxyguanosine dihydroxyacetone phosphate (Figure 2) diethylenetriamine ( 1,4,7-triazaheptane) 2-deoxyguanylyl(5→3)-2-deoxy-5-guanylate thymidine [ 1-(2-deoxy-β -D-ribofuranosyl)thymine] 5-diphosphate thymidine [ 1-(2-deoxy-β -D-ribofuranosyl)thymine] 5-monophosphate thymidine [ 1-(2-deoxy-β -D-ribofuranosyl)thymine] 5-triphosphate dielectric constant (or permittivity) ε-adenosine ( 1,N6 -ethenoadenosine) ε-adenosine 5-monophosphate ( 1,N6 -ethenoadenosine 5-monophosphate) (Figure 12) ethylenediamine ( 1,2-diaminoethane) 9-ethylguanine flavin mononucleotide ( riboflavin 5-phosphate) (Figure 11) glycerol 1-phosphate (Figure 2) guanosine 5-diphosphate guanosine 5-monophosphate guanosine 5-triphosphate hydrogen triphosphate ionic strength inosine 5-diphosphate inosine 5-diphosphate (N1)-oxide inosine 5-monophosphate inosine 5-monophosphate (N1)-oxide (Figure 12) inosine 5-triphosphate general acidity constant general ligand divalent metal ion 1-methylbenzimidazole 9-methyladenine 9-methylguanine 9-methylhypoxanthine methyl thiophosphate 1-methylimidazole nucleobase derivative nucleoside 5-diphosphate

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NMP2 NP Nta3 NTP 4 oh OMP3 oPyN PEEA2 Phen PME2 PMEA2 PMEApp 4 PS2 pUpU3 Py PyN R-DP3 RibMP2 R-MP2 R-PO32 R-TP 4 Tn Tren Tris TuMP2 U UDP3 UMP2 UMPS2 UTP 4 XMP3

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nucleoside 5-monophosphate nucleoside phosphate nitrilotriacetate nucleoside 5-triphosphate octahedral orotidinate 5-monophosphate (Figure 11) ortho-aminopyridine-type ligand 9-[2-(phosphonoethoxy)ethyl]adenine 1,10-phenanthroline (phosphonomethoxy)ethane dianion of 9-[2-(phosphonomethoxy)ethyl]adenine ( Adefovir) (Figure 12) diphosphorylated PMEA2 thiophosphate monoester uridylyl-(5→3)-5-uridylate pyridine pyridine-type ligand diphosphate monoester (Figure 6) D -ribose 5-monophosphate monophosphate monoester (Figure 6) monophosphate monoester and/or phosphonate ligand triphosphate monoester (Figure 6) triethylenediamine ( 1,3-diaminopropane) tris(2-aminoethyl)amine 2-amino-2-(hydroxymethyl)-1,3-propanediol (Figure 14) tubercidin 5-monophosphate ( 7-deaza-AMP2) uridine-type ligand uridine 5-diphosphate uridine 5-monophosphate uridine 5-O-thiomonophosphate (Figure 12) uridine 5-triphosphate xanthosinate 5-monophosphate (XMP – H)3 (X – H • MP)3 (Figure 11)

Species written in the text without a charge do not carry one or represent the species in general (i.e., independent of the protonation degree); which of the two possibilities applies is always clear from the context. In formulas such as M(H;NMP) the H and NMP2 are separated by a semicolon to facilitate reading; yet they appear within the same parentheses to indicate that the proton is at the ligand without defining its location. A formula like (NB H) means that the ligand, here a nucleobase residue, has lost a proton and it is to be read as NB minus H. The term (aq) is used to indicate that water is acting as a ligand.

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Met. Ions Life Sci. 2, 181–240 (2007)

5 Synthetic Models for the Active Sites of Nickel-Containing Enzymes Jarl Ivar van der Vlugt and Franc Meyer Institut für Anorganische Chemie, Georg-August-Universität Göttingen, Tammannstrasse 4, D-37077 Göttingen, Germany

1. INTRODUCTION 2. MODELS FOR COFACTOR F430 3. MODELS FOR SULFUR-RICH NICKEL SITES 3.1. Nickel–Iron Hydrogenase 3.2. Acetyl Coenzyme A Synthase and Carbon Monoxide Dehydrogenase 3.2.1 The C-Cluster 3.2.2 The A-Cluster 3.3. Nickel Superoxide Dismutase 4. MODELS FOR THE UREASE ACTIVE SITE 5. MODELS FOR ACIREDUCTONE REDUCTASE 6. CONCLUDING REMARKS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

1.

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INTRODUCTION

Bioinspired nickel coordination chemistry has flourished in recent years, mainly stimulated by exciting developments in structural biology. Even though the known Metal Ions in Life Sciences, Volume 2 © 2007 John Wiley & Sons, Ltd


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nickel proteins remain rather few in number compared with those containing the more prevalent transition metal ions such as iron, copper, and zinc, their biological functions are truly diverse [1–4]. In fact, each of the well-established biological nickel sites is rather unique with respect to its structure and function. Nickel environments found in biology range from thiolate-rich (CODH and CODH/ACS, Ni-SOD, [Ni,Fe]hydrogenase) to tetrapyrrole-derived scaffolds (F430) in enzymes with redox function, but may also feature histidine-N/carboxylate-O ligation as in ARD or the hydrolytic (non redox-active) urease. Unusual features include, inter alia, a carbamylated lysine (urease) or CO and cyanide ligands ([Ni,Fe]-hydrogenase) or some of the still rare examples of bioorganometallic chemistry (F430, [Ni,Fe]-hydrogenase). In many cases, the functional roles of the unusual ligand environment and details of the overall catalytic mechanism are not well understood. Consequently, the emulation of the fundamental structural and functional motifs of Ni-containing active sites is a fascinating playground for coordination chemists. This is even more true since structural information at atomic resolution has become available recently for many of the important nickel-containing metalloproteins [1], which now provides a solid foundation for investigating their catalytic chemistry and for the design of sophisticated active site models. Synthetic nickel complexes have indeed provided valuable insight into electronic and geometric structures of intermediates proposed for the mechanisms of almost all of the biochemical systems. The biomimetic chemistry of nickel prior to structural elucidation of the metalloproteins (with important contributions to rule out or support hypotheses about active site structures) is summarized in an excellent review by Halcrow and Christou [5]. Essential characteristics of nickel that are pivotal for its chemistry comprise its redox activity with the oxidation states 1 and 3 being accessible under appropriate conditions from the most common state 2, and its Lewis acidity with predominance of coordination numbers five and six [6]. In view of the volume of published work it should be noted that this review does not intend to be exhaustive, and we apologize if our selection does not match the preference of all readers.

2.

MODELS FOR COFACTOR F430

The final step of methanogenesis, i.e., the reaction between methylthioethyl sulfonate (methyl coenzyme M, CH3SCoM) and 7-mercaptoheptanoylthreonine phosphate (coenzyme B, HSCoB) to yield the unsymmetrical disulfide CoM-SS-CoB and methane, is catalyzed by methyl coenzyme M reductase (MCR). The key component of MCR is an unusual yellow Ni tetrahydrocorphinoid cofactor called F430 [7–10] that is found as a noncovalently bound prosthetic group deeply buried in the protein matrix, accessible via a 30-Å-long substrate channel. The macrocycle of F430 is unique and has been termed a hydrocorphinoid because of its high degree of saturation. Structural and spectroscopic characterization as well as the biochemistry of MCR are described in detail by Jaun and Thauer in Met. Ions Life Sci. 2, 181–240 (2007)

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Chapter 8, and mechanistic proposals are only briefly summarized here to provide a background for the overview of synthetic modeling efforts. O

HOOC H HN

CH3

H2N H3C N

N O

COOH

Ni H

N

N COOH

HOOC H O HOOC cofactor F430

Redox chemistry of F430 has been the subject of numerous ESR and MCD studies (both on the enzyme and on model systems) as well as quantum chemical calculations, where NiI, NiII, and NiIII states have been identified and proposed as intermediates in the catalytic cycle [11,12]. Despite extensive spectroscopic work and structural insight, however, the exact mechanism of MCR still remains a mystery. Several principally different mechanisms are currently discussed, and the two most likely ones are sketched in Schemes 1 and 2 [9]. The two mechanisms both involve nickel in the NiI oxidation state as a key player for the reaction, either attacking at the methyl group of CH3SCoM to form a NiIII-CH3 (mechanism I [7]) or attacking the thioether-S to give a covalent NiIISCoM (mechanism II [13,14]). However, mechanism II does not invoke a methylnickel intermediate. Thus, some of the crucial questions are: (i) which features of the ligand architecture of F430 are important for the stabilization of NiI; and (ii) which intermediates play a role in the catalytic reaction (including the question whether a distinct Ni-CH3 is involved). Synthetic model chemistry has significantly contributed to tackle these questions, where {N4} macrocyclic ligands including porphyrins seem to be natural models for the nickel-F430 site in MCR. F430 itself can be separated from the enzyme after denaturation [15,16], and many studies have been performed on the isolated cofactor or its pentamethylester F430M (which shows increased solubility and stability in organic solvents). Lin and Jaun [22,23] have demonstrated that reaction of F430M with Mg(CD3)2 generates a brown, five-coordinate CD3-NiII complex with diagnostic (and paramagnetically shifted) CD3 NMR resonances that are in accordance with a high-spin S 1 species. Reversible reduction of F430M from the NiII form to the NiI form occurs at 0.66 V (in THF), and UV-visible spectra (characteristic absorption maxima at 380 and 750 nm) as well as quasi-axial ESR spectra indicate that reduction is metal based and that the NiI complex is approximately square planar with the unpaired electron in an orbital of d x2–y2 character [17,18]. However, free F430 is Met. Ions Life Sci. 2, 181–240 (2007)

184

VAN DER VLUGT and MEYER CH3-SCoM + H+

N

CH3

N

N

N

N

N

N

NiIII N

NiI H-SCoM CoBS-SCoM

SCoM + H+

CoBS-SCoM

HSCoB N

N

N Ni

II

N

N

CH3

N

NiII N

N

H+

CH4

Scheme 1. Proposed catalytic mechanism of MCR involving nucleophilic attack of NiIF430 at the methyl group of CH3SCoM.

CH3-SCoM CoM S N

N

N NiI

N

CH3

N

CoBS-SCoM

N NiII

N

N

SCoB

HSCoB CH4

CoBS-SCoM

N

N NiII

N

N

Scheme 2. Proposed catalytic mechanism of MCR involving nucleophilic attack of NiIF430 at the thioether-S of CH3SCoM. Met. Ions Life Sci. 2, 181–240 (2007)


185

thermally unstable and the equilibrium mixture contains only 4% of the native form, besides 8% of its 13-epimer and 88% of the 12,13-diepimer. Only F430 (not its epimers) serves as the active site, which has been explained by the stereochemistry at the 12 and 13 positions of the native coenzyme that imposes a planarizing effect on the F430 ring system. Due to its relative inability to ruffle, the core of F430 cannot contract sufficiently to optimally coordinate a small low-spin NiII, but rather favors the larger NiI and high-spin NiII ions [19,20]. This also provides an explanation for the enhanced ligand affinity of F430 in its NiII state with switching from low-spin to the high-spin state concomitant with axial ligand binding. It has been demonstrated that the MCR enzyme is active only if the metal center of coenzyme F430 is in the NiI state [21]. However, while NiIF430M reacts with electrophilic methyl donors such as methyl iodide or methyl tosylate to give methane via a CH3-NiIF430M intermediate, it does not react with the natural substrate CH3SCoM or simple methyl thioethers [17,22,23]. Long before structural information on the active site became available, this led to the proposal of Berkessel [24] and Jaun [25] of a catalytic mechanism in which the addition of a thiyl radical to the S atom of the thioether gives a sulfuranyl radical intermediate in the crucial step (Scheme 3). Such reactivity has been emulated with the model substrate 1, which forms a square planar NiII complex 2. Upon irradiation (i.e., generation of a NiI /thiyl radical pair by excitation of the LMCT band), formation of the spirodisulfide 3 and methane was observed (Scheme 4) [26]. Even if coordination of coenzyme B to the Ni ion in F430 now seems unlikely in view of the crystallographic findings, this model chemistry is important as it represents some similarity to the enzymatic conversion and lends support for the involvement of a thiyl radical that is also implicated in mechanism II shown in Scheme 2. CoB

CH4

S

N

N NiII

N

HSCoB

N

CoB N

CH3

N NiI

NiII N

S

N

N N

N CoB CoM

S S

CH3

N

N NiI

CoMS-SCoB N

N

CH3-SCoM

N

Scheme 3. Proposed catalytic mechanism of MCR involving addition of a thiyl radical to the S atom of CH3SCoM. Met. Ions Life Sci. 2, 181–240 (2007)

186


H3C S

S

H3C S

NiII S

S

CH3

S

dark

S NiI S

CH3

2

SH

S CH3

–CH4

1 CH3

(L) NiII S

S CH3 H3C S

+

S

NiI S

S

S CH3

S 3

Scheme 4.

The reductive dealkylation of 4 to afford 5 (Scheme 5) was noted by Sellmann et al. [27] and studied mechanistically by the Kovacs group [28]. NiI intermediates have been detected by ESR spectroscopy, and formation of the organic products (C2H4 and ethylene sulfide) was explained by transfer of the NiI d x2–y2 unpaired electron to a C–S σ*-orbital, inducing homolytic C–S cleavage. A somewhat related S→NiIII electron transfer was proposed by Berkessel for the oxidative cleavage of the -CH2SPh pendant arm from a series of substituted Ni(dihydrosalen) derivatives after metal-centered oxidation [24,29]. The study of NiI with porphyrin-type {N4} ligands has received a major impetus from the unique redox chemistry of F430, and nickel tetrapyrroles [30], including core-modified porphyrins, [31] have been studied extensively in this context. Major factors that determine the NiII /NiI redox properties are the core size, the flexibility of the macrocycle, and the electronic properties of substituents [32,33]. NiII has a (dz2)2 electronic configuration and requires short Ni–N distances, forcing the porphyrin macrocycle to adopt a nonplanar conformation. Switching from low-spin to high-spin NiII (e.g., by axial ligation or photoexcitation) induces a considerable change in the conformation such as a flattening of the macrocycle. Met. Ions Life Sci. 2, 181–240 (2007)


187

S S S

2–

NiII

S

+ 2e

S

S

–C2H4

S

NiII

S S

S –

5

4

S

+e

S S S

NiI

+e

S

–

S

S

S

S

NiII

S

–

S

S S

–

–C2H4

S

NiII

–

S S

Scheme 5.

As a consequence, porphyrins with bulky meso alkyl substituents that enforce nonplanar deformations lead to a decrease in binding affinity for axial ligands, or binding is even completely inhibited for the more nonplanar systems [34]. The reduction of regular NiII porphyrin systems very often results in the formation of a ligand-based radical rather than in metal-based reduction. Molecular mechanics calculations have been used to examine the effect of macrocyclic reduction on the core-hole size of porphyrins, and findings have been applied to the nickeltetrapyrrole F430 cofactor [35]. Macrocycle reduction at the two methine protons provides the F430 macrocycle with the flexibility necessary to accommodate both high-spin (six-coordinate) and low-spin (square planar four-coordinate) NiII [19]. Complexes of isobacteriochlorins, in particular octaethylisobacteriochlorin (oeibc), are frequently used as F430 mimics because they can be reduced to isolable NiI complexes with ESR spectral parameters similar to those of reduced F430 or F430M (g 2.20, g⊥ 2.07, AN 9.8 G) [36,37,38,39,40]. [NiI(oeibc)] (6) has been studied by EXAFS, ESR, and ENDOR spectroscopy, and bond lengths and spectroscopic data were correlated [41]. X-ray absorption near-edge data show that at low temperatures 6 binds axial ligands to form five-coordinate complexes [42]. However, binding of pyridine does not cause the axial-to-rhombic ESR spectral changes that have been observed for F430. The stability of [NiI(oeibc)] is greatest in polar solvents. With weak acids such as water, alcohols, and thiols it affords H2 with an apparent stoichiometry of 2:1 NiI:H2 [43]. The reactions of [NiI(oeibc)] with alkyl and aryl halides give organic products that result from reduction or dehydrohalogenation [43,44]; alkene formation is favored by polar solvents, and isomerization of alkenes occurs during reaction. This implies the intermediacy of Ni-H and Ni-R species in the reactions. With CH3I or Met. Ions Life Sci. 2, 181–240 (2007)

188


methyl p-toluenesulfonate 6 reacts to give neutral [NiII(oeibc)] in quantitative yield as well as CH4 and I (or p-toluenesulfonate, respectively) in lesser yields. Reactivity patterns for different alkyl halides RX are consistent with the rate-limiting step being SN2-type nucleophilic attack at RX by [NiI(oeibc)] to generate a transient [R-NiIII(oeibc)] species, in analogy with the postulated reaction of NiIF430 to form CH3-NiIIIF430 according to mechanism I (Scheme 1). Subsequent decay of [RNiIII(oeibc)] is complex [45]. For investigating the reaction of NiI and NiII porphyrin anions with alkyl radicals R the pulse radiolytic technique has been applied [46]. Reactions take place readily, but the resulting R-NiII or R-NiIII species are unstable and decay within seconds or milliseconds, respectively. Although the NiI-complex of oeibc is very resistant to further reduction, in the presence of proton donors reduction of the macrocycle ring occurs [39], affording initially the NiI-complex of 7, the first complex of this precorrin to be identified. −

Et

Et Et

Et Et

N

Et H

N

H Et

H

N

N

N

N

Et H

NiI N

N

Et

Et Et

Et 6

Et

Et Et

H H

Et

7

Since the catalytic cycle of F430 is postulated to involve reduction of the NiII resting state to NiI followed by formation of a methyl-NiIII transient from which methane is formed, the question arose whether the tetrapyrrole ligand of F430, which is unusual in its ability to stabilize NiI, can also give rise to high-valent NiIII. DFT calculations suggest that NiIII is indeed accessible, the special architecture of F430 allowing occupation of the dx2–y2 orbital [47]. Interestingly, the electronic spin density profiles of this (dx2–y2)1 NiIII species can be deceptive and closely mimic NiI species. The putative NiIII species in F430 has also prompted considerable interest in high-valent Ni-porphyrin compounds [48,49]. Some useful models, both electrochemically as well as functionally, are the Ni-complexes with cyclam-derived macrocyclic tetraaza ligands such as 8–10 [50–52]. A set of NiI complexes [53] of these tetraaza ligands has been structurally characterized [54–57]. Reaction of [CH3Co(dmgBF2)2 L] (dmgBF2 (difluoroboryl)dimethylglyoximato, L py, PEt3) with two equivalents of [Ni(tmc)] gave [Co(dmgBF2)2L] , [Ni(tmc)] 2 and [Ni(tmc)CH3] in 80% yield [57]. R,R,S,S-[Ni(tmc)CH3](BAr4') (BAr4' B(3,5-(CF3)2C6H3)4 ) was characterized by X-ray diffraction (Figure 1) [57]. The products and stoichiometry of the reaction are consistent with a three-step mechanism initiated by electron transfer from [Ni(tmc)] to

Met. Ions Life Sci. 2, 181–240 (2007)


Figure 1.

189

Molecular structure of [NiII (tmc)CH3] [57].

[CH3Co(dmgBF2)2L] . The second step is rapid CH3-Co bond homolysis yielding [Co(dmgBF2)2L] and · CH3, and then the · CH3 radical is captured by the second equivalent of [Ni(tmc)], yielding [Ni(tmc)CH3]. The NiII complexes were found to be homogeneous catalysts for the dehalogenation of cyclohexyl bromide by sodium borohydride, with the effectiveness depending on the solvent and the structure of the complex. [NiII(tmtaa)] (8) is ESR-silent as expected for a square planar d8 system, while the ESR spectrum of a degassed mixture of 8 and NaBH4 shows a rhombic signal (g储 2.26 and g⊥ 2.13) in accordance with a nickel(I) species [50]. This signal rapidly decreases after addition of cyclohexyl bromide. The greater anisotropy of the ESR signal compared with that of authentic [NiI(tmtaa)] (produced by sodium amalgam reduction) suggests I the coordination of either hydride or BH 4 to Ni [50,52]. None of these complexes, however, cleaves CH3SCoM to methane. It should be noted that nickel boride (a heterogeneous catalyst for the dehalogenation of cyclohexyl bromide) produces a small amount of methane from CH3SCoM and NaBH4, but preferentially cleaves the CH2–S bond of the substrate [52]. Irradiation of solutions of 11 and 0.01 M HCO2Na with γ -rays or 3 MeV electrons afforded the corresponding NiI-species. Injection of CH3-SCoM gave methane, blank experiments without the Ni complexes did not [58]. [NiII(tmc)] 2 (tmc tetramethylcyclam) binds CH3-SCoM readily, but coordination seems to occur via the sulfonate end group [63]. 2+

N

N

N N

8

N

H N

N

N

N

H

10

N

Me

Ni N

H

9

Me

Ni N

H

2+

2+

H

Ni

Ni N

N

H

Me

N

N

Me

11

Met. Ions Life Sci. 2, 181–240 (2007)

190


If alkyl radicals generated by pulse radiolysis are reacted with [NiII(tmc)]2, short-lived R-NiIII species with optical absorption bands similar to those of other NiIII-complexes are formed (300, 350, and 520 nm) [59,60]. These species rapidly decompose via various pathways involving Ni–C bond homolysis. R-NiIII compounds apparently are thermodynamically less robust than the corresponding R-NiII analogs. The cyclam-based complex 12 bearing a methylthio pendant arm has been proposed as a model for F430 [61]. While participation of the thioether group in Ni coordination is observed neither in solution nor in the solid state, the NiII species is reversibly reduced to NiI around 0.7 V compared with SCE, followed by thioether bond cleavage and formation of a thiol group at more negative potentials. Reduction of 12 with Na/Hg in DMF produces small amounts of methane. In related cyclam-derived NiII systems with one, two or four methylthio-substituted pendant aliphatic chains, coordination of one of the thioether-S to give fivecoordinate complexes is observed (e.g., in 13; Figure 2) [62]. NiII is reversibly reduced to NiI between 0.64 and 0.77 V compared with SCE. At more negative potentials the thioether is cleaved, forming a thiol. MeS

2+

2+

MeS MeS

N

N Ni

N

N

N Ni

N 12

N

N

13

Pentacoordinate thiophenolate complexes of NiII complexes with cyclamderived ligands have been independently prepared and structurally characterized, e.g., in [NiII(tmc)(SPh)] [63] (Figure 3). This mimics the situation of the S-bound CoM as postulated in mechanism II (Scheme 2) and the binding motif found in the active site in the MCRox1/silent state [7].

Figure 2.

Molecular structure of the cationic portion of 13 [62].

Met. Ions Life Sci. 2, 181–240 (2007)


191

Figure 3. Molecular structure of [NiII (tmc)(SPh)] [63].

3. 3.1.

MODELS FOR SULFUR-RICH NICKEL SITES Nickel—Iron Hydrogenase

Spurred by a combination of academic curiosity-driven interest in gaining further understanding of the bio-(inorganic) chemistry involved in these metalloenzymes and the promise of developing cheap H2-producing catalysts, the enzyme class of the hydrogenases, which catalyze the interconversion of protons and H2, has been the subject of intense research, not solely in the field of biochemistry, but also in the inorganic chemistry community [64]. H2 2 H 2 e H 2 O D2 HDO HD Nowadays, three main types of hydrogenases are distinguished (see also Chapter 7): [Fe]-only hydrogenases, [NiFe]-hydrogenases, and the Fe–S clusterfree hydrogenases. Despite structural similarities between the active sites in the first two types of hydrogenases, there appears to be no genetic evolutionary relation. Within the family of [NiFe]-hydrogenases (Figure 4), four subclasses are defined: H2-uptake, H2-evolving, bidirectional H2-activation or H2-sensing enzymes. To date, most research has focused on the first class, and this is at least in part because the available X-ray crystallographic data deal exclusively with this subclass [65,66]. Besides the uptake of H2 by these enzymes, [NiFe]-hydrogenases have also been shown to catalyze H/D exchange, without the use of electron donors or Met. Ions Life Sci. 2, 181–240 (2007)

192 (a)

VAN DER VLUGT and MEYER (b)

Fe S S Fe L S S S Fe S Fe Fe CO NC Fe S CN OC C O

(c) Cys

OC

Cys

Cys S S

NC Fe NC

Ni

S Cys S Cys

Cys S S

OC Ni NC Fe S Cys O NC S H Cys

Figure 4. Structures of the active sites found in: (a) [Fe]-only hydrogenase; (b) reduced state, [NiFe]-hydrogenase; (c) oxidized state, [NiFe]-hydrogenase.

acceptors. To date, accurate structural and functional modelling of the [Fe]-only hydrogenases has been of key interest, and in recent years a number of research groups have made successful efforts and significant contributions, which will not be discussed here; the interested reader is referred to the recent literature [67,68,69]. The active site of [NiFe]-hydrogenase contains a sulfur-only coordinated Ni center together with an Fe center with two terminal CN ligands and one terminal CO ligand. When the system is in the oxidized state, crystallographic analyses point to the presence of a third bridging ligand between the Ni and Fe centers, most probably oxygen-based. Three major unique structural features stand out for this active site: (i) the atypical non square planar, tetrahedral geometry around the sulfur-only coordinated nickel atom; (ii) the presence of two cysteine-derived thiolate bridgehead groups; and (iii) the trigonal bipyramidal Fe-center which contains the highly toxic and previously bio-irrelevant ligands CO and CN. The catalytic mechanism of the [NiFe]-hydrogenase active site is still not fully resolved, despite intense investigations from a number of specialized groups, using a variety of spectroscopic measurements, including ESR, X-ray absorption, IR, EXAFS, Mössbauer, ENDOR, and UV/Vis spectroscopy, and complemented with computational chemistry. Several states for the [NiFe]-hydrogenase active site are now distinguished, which indicate a rich redox chemistry for the active site and electronic communication between the two metal centers, i.e., both metals are influenced by the redox reactions. We will shortly introduce the most important features of the proposed mechanism and the corresponding states of the [NiFe]-hydrogenase active site, to allow a more comprehensive relation to the model systems described in this chapter. For a full account of these results and the derived catalytic mechanism, the reader is referred to Chapter 7 of this volume. Purification of [NiFe]-hydrogenase in air results in an oxidized protein product that exhibits two ESR signals, termed Ni-A and Ni-B, both inactive. The precise nature of the structural difference between these two states is still unclear. X-ray diffraction and several spectroscopic measurements indicate the presence of a third oxygen-based bridging ligand, which for Ni-B is most likely a hydroxide. However, Ni-B can be readily reduced anaerobically by H2 or dithionite in a Met. Ions Life Sci. 2, 181–240 (2007)


193

two-electron process, yielding active enzyme in the so-called Ni-C state, whereas Ni-A is only slowly converted to that same species. All three states contain paramagnetic NiIII, as derived from their respective ESR spectra. Ni-A, Ni-B, and NiC have also been termed the ‘inactive’, ‘ready’ and ‘active’ state of the enzyme’s active site. The oxygen-containing ligand is believed to inhibit any catalytic activity in Ni-A and Ni-B; no oxygen-bridging ligands are found in the crystal structures obtained for the active enzyme. One-electron reduction of the Ni-A and Ni-B states yields ESR-silent NiII intermediates termed Ni-SU and Ni-SI; Furthermore, one-electron reduction of Ni-C yields Ni-R (SU silent unready, SI silent ready, R reduced active) where Ni-R is again ESR-silent. The redox potentials for the NiIII /NiII couples found in the family of [NiFe]-hydrogenases fall in the range of approximately 0.39 to 0.64 V compared with SCE. This is in marked contrast with the more positive values (between 0.50 and 1.50 V compared with SCE) for most synthetic Ni-based complexes. Both Ni-SI and Ni-R can take up a proton. Recently, ENDOR and HYSCORE spectroscopy have provided evidence for the presence of a hydride bridging the Fe and Ni centers [70–72]. The influence of the thiolate ligands during protonolysis or reaction with H2 is still unclear. The vibrational band of the CO ligand on the Fe center, which remains in the FeII state throughout all detectable states, as observed by ENDOR and ESR spectroscopy, falls in the range 1914–1952 cm1 in the various states. These small fluctuations are further indication that no FeII /FeIII transitions occur during catalysis. Modeling of the [NiFe]-hydrogenase active site characteristics has focused for a long time on simple mononuclear complexes. Recently, a comprehensive review appeared covering relevant mononuclear nickel complexes with a variety of ligands as well as important Fe(CO)x(CN)y species [73]. We will discuss a few noteworthy model systems that show significant functional or structural resemblance with the native enzyme. The group of DuBois has described mononuclear Ni-complex 14, containing two Et2PCH2N(Me)CH2PEt2-ligands, as a functional model for [NiFe]-hydrogenase activity, in that it is able to produce H2 by electrochemical reduction of protons. Furthermore, H2 is heterolytically cleaved by this complex [74,75]. H N

2+

N

H H P P Ni P P

H P N

N

P

Ni

2+

P P

14

Inspired by a possible participation of the thiolate ligands during H2 uptake or protonolysis of different states of the active site, Liaw et al. [76,77] reported on the interaction of a thiol with a mononuclear Ni-complex. To this end they reacted the tetradentate ligand P(o-C6H4SH)3 with a NiII salt to obtain complex 15, which Met. Ions Life Sci. 2, 181–240 (2007)

194


showed an interaction of the SH proton with both the nickel ion and a coordinated thiolate group, as observed by X-ray crystallography, IR and NMR spectroscopy [76,77].

H

SH

SH

[Ni(ER)3CO]–

RE

P

S

NiII

S

S P

HS 15 ER = SePh, SPh

The group of Sellmann has reported on H2 /D exchange at a mononuclear distorted tetrahedral Ni-complex 16, ligated by a thioether, two thiolates, and one iminophosphorane [78]. Such H/D exchange, which is presumed to involve heterolytic H2 cleavage, is used in the biochemical assaying of hydrogenase activity [79]. The proposed mechanism involves protonation of one of the thiolate moieties, after which scrambling and production of HD takes place; this result indicates that H2 heterolysis can in principle occur at a redox-silent NiII center and does not require a NiII to NiI reduction step.

S 16

S S Ni

+ D2

NH PPr3

D S Ni S

D S NH PPr3

–HD

S Ni S

S ND PPr3

More recently, the heterobimetallic nature of the dinuclear center, which clearly creates many challenges for the coordination chemist, has been explored in small molecule models. We will further concentrate here on modeling efforts that truly incorporate both metal atoms. An important synthetic route to thiolate-bridged heterobimetallic complexes is the addition of a metal complex with labile ligands to a mononuclear metalthiolate species; however, the high tendency of metal-thiolate complexes to aggregate, forming undesirable homometallic clusters, severely limits this strategy. In order to suppress cluster formation, thiolate-containing polychelating ligands are frequently employed. The chelate effect then prevents dissociation of the thiolatecontaining ligand and directs subsequent reactions with metal salt and complex reagents towards the terminal thiolate donors. Mixed NiFe complexes have most commonly been obtained via reactions of NiII-thiolate complexes with labile Fe species, giving a range of synthetic models for the [NiFe]-hydrogenase active site. Met. Ions Life Sci. 2, 181–240 (2007)


195

A number of thiolate-bridged NiFe systems have been reported to result from N,N-bis(2-mercaptoethyl)-1,5-diazacyclooctane (H2L), which reacts with NiII to first yield 17 as a suitable precursor complex. This has subsequently been treated with a variety of FeII compounds [80]. The reaction of 17 with Fe2 (CO) 9 leads to Ni(L)Fe(CO) 4 (18). In this case the Ni and Fe centers remain in their original oxidation states of NiII and Fe0, respectively, and the Fe(CO) 4 fragment is simply bound to Ni(L) via a single thiolate bridge at a large Ni–Fe distance of 3.76(1) Å. This NiFe complex was one of the early examples that confirmed the ability of Ni-coordinated thiolates to bind to Fe-carbonyl fragments with some control over stoichiometry. Upon oxidation with NO(BF4), complex 19 is formed, in which Fe0 has been oxidized to FeII. The FeII ion in 19 links two Ni(L) units each through two thiolate bridges. The Ni ions remain in the NiII formal oxidation state in a square planar N2S2 donor set, and the Ni–S distances do not change significantly. The Ni–Fe distance decreases from 3.76(1) Å in 18 to about 3.09 Å in 19, accompanied by a decrease in the Ni–S–Fe angle from 89.3(4) to 76.99(7). In the presence of cobaltocene as a reductant, 19 reacts with CO to regenerate 18. CO CO CO N Fe N Ni S CO S

N N Ni S S

Fe2(CO)9

17

S

N

OC

Ni

NO(BF4)

N

CoCp2, CO

19

18

CO

2+

Fe S

S S Ni N N

When, instead of N,N-bis(2-mercaptoethyl)-1,5-diazacyclooctane, the more flexible metalloligand Ni(dsdm) (H2dsdm N,N-dimethyl-N,N-bis(2-sulfanylethyl) ethylenediamine) was reacted with Fe2 (CO) 9, the unexpected trinuclear 48-electron NiFe2 complex 20 was obtained [81]. Similar results were also found by the group of Schröder with a bisthiolate-bisthioether ligand [82]. For the NiFe2 complex 21 with this latter ligand, the resulting species obtained after one-electron reduction showed ESR hyperfine coupling when enriched with 61Ni, and an average shift of 70 cm1 for the CO bands in the IR spectrum.

N

S

N

OC S OC Fe OC

S CO Fe CO CO 20

S Ni

Ni OC OC OC

S Fe 21

S CO Fe CO CO

Another unusual structural motif with remote analogy to the [NiFe]-hydrogenase active site was reported by Schröder et al. [83]. Reaction of the Ni(tsalen)-complex Met. Ions Life Sci. 2, 181–240 (2007)

196


(tsalen N,N-ethylenebis(thiosalicylideneiminato)) with Fe2(CO) 9 yielded the dinuclear species 22, together with the trinuclear derivative 23.

N

S

N

Fe 2(CO)9

Ni

Ni N

S

N

S

Ni(tsalen)

CO Fe CO CO

N Ni

+

S

S

OC OC Fe OC

S

N

22

CO Fe CO CO

23

Reaction of pentadentate N,N-dimethyl-N,N-bis(2-mercaptoethyl)-bis(aminoethyl)sulfide (H2L) with NiII yielded the corresponding metalloligand Ni(L). When treated with FeCp(CO)2I, this NiII-thiolate complex gave rise to [Ni(L)FeCp]I (24), which is related to the reduced form of the [NiFe]-hydrogenase enzyme [84]. The thioether donor occupies the apical position of a square pyramidal coordination geometry, and the Ni–S(thioether) bond (2.283(6) Å) is shorter than the Ni–S(thiolate bridge) distances (2.308(6) and 2.315(6) Å). The Ni( µ-S)2Fe core exhibits a very acute Ni–S–Fe bridging angle of 58–59 (compared with 74 in [NiFe]-hydrogenase from D. gigas) [85,86], which brings the two metal ions within close proximity (2.54 Å, i.e., similar to the value of 2.5–2.6 Å observed for the Ni–Fe distance in the Ni-SI form of the enzyme) [87].

N S

Ni

Ph2 P

S S

Fe

N 24 N Ph2 Cl P Ni S Fe S CO S P CO Ph2 26

CO

Ni S Fe CO CO S P Ph2 25 N S S Fe S CO

OC S Ni

Fe S N S

27

Instead of terminal N-donor ligands at the Ni center, soft phosphines have frequently been employed since these are better suited to emulate the soft cysteine sulfurs of the enzyme. For instance, addition of Fe3 (CO)12 to (dppe)Ni(pdt) (dppe l,2-bis(diphenylphosphino)ethane, pdt 1,3-propanedithiolate) gives Met. Ions Life Sci. 2, 181–240 (2007)


197

the neutral heterobimetallic complex (dppe)Ni(pdt)Fe(CO)3 (25) [84]. It should be noted that the same 1,3-propanedithiolate unit is thought to be relevant in the active site of [Fe]-only hydrogenase [88,89]. The Ni coordination sphere has a distorted tetrahedral arrangement and the angle between the Ni–S1–S2 and Fe–S1–S2 planes of 80.3 compares well with an angle of 81 in the enzyme. Several bis(thiolato) bridged NiFe-complexes have been prepared by the reaction of [Fe(L)(CO)] [L3 N(CH2CH2S)33] and appropriate NiII species. Treatment with [(dppe)NiCl2] under CO gave [{Fe(L)(CO)2-S,S}NiCl(dppe)] (26) where the NiII is found in a distorted square pyramidal geometry with two bridging thiolate moieties occupying two cis positions in the basal plane [90]. Also coordinated to this Ni center are a dppe ligand in the basal plane and a Cl ligand in the axial position. This particular complex was the first literature example of a model complex featuring a bis-thiolate-bridged heterodinuclear Ni–Fe core with CO coordinated to the FeII center. When the reaction was performed under nitrogen, the monocarbonyl rather than the dicarbonyl complex was formed. Reaction of [Fe(L)(CO)] with the DMSO-solvate of NiCl2 gave a trinuclear FeNiFe complex 27 in which the central NiII ion resides in an almost regular tetrahedral thiolate environment [91]. Sellmann et al. have reported on a series of NiFe complexes, derived from the {S32} ligand bis(2-mercaptophenyl)sulfide. Complex 29 resulted from the reaction of dinuclear 28 with labile [Ni(bdt)(PMe3)2] (bdt 1,2-benzenedithiolate) [92]. Its bridging Ni–S distances (2.2302.241 Å) are distinctly longer than the terminal Ni–S distances (2.142.25 Å), but the latter are typical for diamagnetic four-coordinate NiII-thiolate species. The [Ni(SR)2] fragment withdraws electron density from the Fe center of the [Fe(CO)L2 (SR)2] core, leading to a blue-shift of the ν(CO) frequency.

O C

S S OC

Fe

S S

Fe S

C O

28

S

CO S

S

Ni

60˚C, 3h

PMe3

PMe3 PMe3

S S

Ni

S

Fe S

THF

PMe3 CO

S

29

In related work the trinuclear cluster (30) was obtained in 19% yield from the same reaction of [(Fe(CO)2 ({S3})2] with [Ni(bdt)(PMe3)2] [93]. The Ni–Fe distances are ⬃2.6 Å, while the Fe–Fe distance is 2.4 Å. Despite its trinuclear nature, this complex shows properties that allow for moderately good structural resemblance and, which is rarely observed, also provides for a functional model, since it was shown to reduce protons to yield H2 [93]. Cyclic voltammetry revealed Met. Ions Life Sci. 2, 181–240 (2007)

198


+

Me3P

S

S PMe 3 Ni Ni S S S Fe S C O

30

[FeCp2]PF6

Me3P

S

S PMe 3 Ni Ni S S S Fe S C O

31

quasi-reversible reduction phenomena at 1071 mV, 95 mV and 844 mV compared with NHE, with the second redox couple falling within the potential range observed for typical [NiFe]-hydrogenase redox reactions. Upon treatment of complex 30 with HBF4, the CO vibrational IR band shifted from 1916 to 1976 cm1, similar as observed for the stoichiometric (one equiv.) chemical oxidation of complex 30 with [Fe(Cp)2]PF6. This indicates that a one-electron transfer reaction takes place. The chemically oxidized species 31 proved stable in solution for several weeks and its molecular structure (Figure 5) was very similar to that obtained for 30. In the ESR spectrum a rhombic signal was observed with g values around 2. Furthermore, by 1H NMR spectroscopy, the presence of free H2 could be observed after addition of HBF4 to complex 30, while the reversal of the oxidation was easily achieved by addition of (NBu4)BH4.

Figure 5. Molecular structure of the cation of complex 31 [93]. Met. Ions Life Sci. 2, 181–240 (2007)


Figure 6.

199

Molecular structure of 33 [94].

Tatsumi et al. [94] very recently obtained another close structural model, via an elegant synthetic route that started from (PPh4)[Fe(CO)3 (CN)2 (Br)]. The intermediate 32 was then reacted with a dithiocarbamato-nickel species to obtain the diamagnetic complex 33 (Figure 6). Its molecular structure and spectroscopic features are consistent with low-spin NiII and FeII ions. −

N C OC OC

Fe

Br CO

C N

K2pdt

[Fe(CO)2(CN)2(pdt)K]−

Ph3P

−

S

Br Ni

S

NEt2

NC OC OC

32

S S Fe

Ni CN 33

S S

NEt2

For this complex, the IR spectroscopic data show vibrational bands at νCN 2110 and 2094 cm1, while the Raman spectrum shows only one CN stretching band for νCN at 2113 cm1. These combined data indicate a mutual trans coordination of the two CN ligands. This is in accordance with the X-ray crystallographic data, where the Fe–C(N) and Fe–C(O) distances differ substantially. However, the IR region for the CO ligands showed four bands at various intensities. The nickel center is slightly distorted square planar. The Ni–Fe distance of ⬃3.05 Å corresponds well with those found in two native enzymes (D. gigas and D. fructosovorans) at 2.9 Å, despite the fact that in the oxidized state in D. gigas a third bridging ligand is present. The octahedral geometry around the Fe center (as well as coordination of a second CO ligand) does not exactly match the native system. Nonetheless, the stepwise preparation of heterodinuclear complexes starting from suitable Fe precursors seems promising for future attempts to mimic the intriguing substitution pattern of the [NiFe]-hydrogenase active site. Met. Ions Life Sci. 2, 181–240 (2007)

200


3.2. Acetyl Coenzyme A Synthase and Carbon Monoxide Dehydrogenase 3.2.1.

The C-Cluster

Unifunctional carbon monoxide dehydrogenase enzymes catalyze the conversion of CO to CO2, which occurs at an active site containing a biologically unique NiFe4S5 cluster, as was unequivocally deduced from crystallographic studies (see also Chapter 9). (CODH)

CO2 + 2 e– + 2 H+

CO + H2O

In fact, three structurally related but distinctly different crystal structures have been determined for the CODH active site (Figure 7), obtained from three different enzyme sources [95–97]. Before the elucidation of the general features of the active site of this enzyme, the so-called C-cluster, most modelling attempts were guided by spectroscopic data obtained from studies with the native enzyme and directed to obtaining NiFe3S4 and NiFe4S4 systems, wherein the nickel was bound to the FeS cluster through either one or two thiolate bridging ligands. Although the catalytic mechanism for CODH is still unresolved, a proposed scheme involves coordination of H2O to the exogenous FeII center, which is deprotonated to yield a hydroxide ligand. CO coordination to the NiII center is then followed by nucleophilic attack from the -OH onto the bound CO. The carboxylic acid fragment is deprotonated, CO2 is released upon transport of two electrons (most likely from the Fe3S3-cluster) and the resulting Ni0 center is reoxidized to NiII [98]. The actual structural data for the native system have been known for several years now, yet the most faithful models for the unprecedented Ni-containing FeScubane cluster remain those synthesized by Holm et al. [99] well before the crystallographic analyses appeared. These cuboidal mixed-metal systems have been obtained by reaction of linear trinuclear (NEt4)3[Fe3S4 (SEt) 4] with Ni(PPh3) 4, (a)

(b)

SCys

S

Fe S

Ni

S Fe Fe SCys

SCys

(c) X

SCys Fe

S S SCys

Fe

SCys NHis

S

S S

Ni

SCys

Fe S Fe SCys Fe SCys NHis

SCys

CO

SCys Fe S

S

Ni

S Fe Fe

SCys

S SCys

SCys Fe SCys NHis

Figure 7. Structures of CODH C-cluster active sites, as determined by X-ray crystallography: (a) reduced, active form in C. hydrogenoformans [95]; (b) non-reduced, COpreincubated form in R. rubrum [96], X modeled as CO; (c) from bifunctional CODH/ ACS in M. thermoacetica [97]. Met. Ions Life Sci. 2, 181–240 (2007)


201

which led to a reductive rearrangement to give the stable cubic species (34), with the four-coordinate nickel center incorporated into this fragment and one PPh3 remaining [99].

S

EtS

Fe

EtS

Fe S

S S

–

Ph3P

3–

Ni

SEt

SEt

S Fe

S

Ni(PPh3)4

Fe

Fe

S

SEt

SEt

S

Fe

34

EtS

Alternatively, the incomplete cubic Fe3S4 cluster 35, featuring a tridentate trithiolate ligand, was reacted with NiCl(PPh3)3 to give the same structural core [100]. Besides the absence of the non-cuboidal exo-Fe-center, another inconsistency compared with the natural system is the tetrahedral geometry around the paramagnetic nickel center. Recently, a subtle, but significant improvement in structurally modeling the C-cluster has been achieved, by reaction of 36 with the strongly chelating ligand dmpe, leading to a more accurate low-spin square planar

2–

S

S

S

Fe

S

S

Fe

S Fe

S

Ni(PPh3)4

Fe

S

Ni

Fe

S S

2–

Ph3P

S S

Fe

S

S

35

36

dmpe

S

S

S

S

S S

S

S

S

= S

S

Me2P

PMe2

2–

S

Ni S

S

S Fe

Fe

S Fe

S S

S

37

Met. Ions Life Sci. 2, 181–240 (2007)

202

Figure 8.


Molecular structure of the anion portion of 37 [101,102].

geometry around the NiII ion. During this reaction, one Ni(µ3-S) bond is broken to accommodate the diphosphine ligand, concomitant with a structural rearrangement to square planarity around the nickel center, yielding species 37 [101,102] (Figure 8). Although there are no experimental procedures reported to date for the incorporation of the exo-Fe site, and despite the presence of the hardly biological phosphine donor groups, this tetranuclear NiFe3S4 complex provides the most accurate structural mimic for the CODH active site from the unifunctional enzyme C. hydrogenoformans to date. Heterobimetallic functional models for the catalytic function of the CODH enzyme are still absent, to the best of our knowledge; although a number of NiFe complexes have been prepared (see the previous section on [NiFe]-hydrogenase) no reports on any (attempted) conversion of CO to CO2 have appeared. However, dinuclear complexes with the general structure 38 have been shown to be active Met. Ions Life Sci. 2, 181–240 (2007)


203

catalysts for CO oxidation at room temperature in aqueous medium with electron transfer to methylviologen and production of CO2 [103]. R1 R2

R3

N N

Ni O

S

R2

R2

S

O Ni

R3

N

N R2

R1 38

3.2.2.

The A-Cluster

Bifunctional CODH/acetyl coenzyme A synthase (ACS) enzymes are found in both acetogenic bacteria and methanogenic archaea. In the former, they catalyze the conversion of either CO or CO2 to acetyl-CoA, which is closely related to acetic acid. This also explains part of the interest in the mechanism and modeling of this enzyme’s active site, while the present-day industrial production of acetic acid from methanol and CO utilizes expensive Rh or Ir as catalyst instead of Ni and Fe. (ACS)

CO + CH3-CoIII-CFeSP + CoASH

O H3C C SCoA + CoI-CFeSP + H+

Crystallographic details on the active site of ACS (the A-cluster, Figure 9) were reported by two independent research groups [104,97]. It consists of an Fe4S4 cluster unit, which most likely functions as electron transfer unit, linked to a tetrahedrally coordinated proximal Ni atom (Nip) and a square planar distal Ni center (Nid). The former shows coordination to three thiolate ligands and a still unknown fourth small molecule ligand, while the latter is coordinated by an N2S2

O CysS

S

Fe S

Fe

S

Fe CysS

Figure 9.

Fe

S N SCys Nip Nid L N S

S SCys

O

O

Structure of the A-cluster active site of acetyl coenzyme A synthase (ACS). Met. Ions Life Sci. 2, 181–240 (2007)

204


ligand with a peptide backbone provided by a cysteine-glycine-cysteine (CysGly-Cys) sequence. The catalytic mechanism of ACS is still not fully clarified, but a combination of spectroscopic measurements, including ESR, X-ray absorption, IR and EXAFS spectroscopy, have provided several clues. We will briefly introduce the most important features of the proposed mechanism and the corresponding states of the ACS active site, to allow a more comprehensive relation to the model systems described in this chapter. For a full account of these results and the derived catalytic mechanism, the reader is referred to Chapter 9. In fact, two distinctly different mechanisms have been proposed; the first contains a Ni0 center as the proximal site, at which CO coordinates, followed by methyl-group transfer from a methylated corrinoid–iron–sulfur protein, to generate a NiII(CO)(CH3)-fragment. CO insertion (or methyl migration) produces an acetyl group which is attacked by deprotonated CoA to yield acetyl-CoA as the product. In this mechanism, the distal Ni site and the Fe4S4 cluster play no direct functional role in substrate binding [105,106]. Another mechanism starts from a proximal NiII center, which is reduced to a NiI species by electron transfer. CO association and methyl-group transfer yield a NiIII species. One-electron reduction gives the analogous NiII center, which then undergoes CO insertion and dissociation of acetyl-CoA. This mechanism is not balanced with respect to electron count [107]. There are several very demanding requirements to be fulfilled in order for a small molecule system to be an accurate structural model for the ACS active site: (i) a square planar redox-inactive NiII ion, coordinated to two carboxamido-N and two thiolato-S atoms, mimicking the diamagnetic d8 Nid site; (ii) a NiII complex in square planar or tetrahedral geometry (Cu or Zn analogues are optional) ligated by two sulfur donating ligands, a bridging thiolate connected to an Fe4S4 cluster or model thereof and a fourth ligand with proven coordinative lability, to model the Nip site; (iii) reduction of the NiIIp-center to NiI concomitant with CO coordination; the resulting ESR signal should have g values around 2; (iv) the Nip site should be coordinatively unsaturated, allowing for CO coordination and subsequent insertion into a Ni-CH3 bond; (v) the Nip site should exhibit labile binding, and treatment with (excess) chelator, e.g., phenanthroline, should result in demetallation [108,109]. Before the crystallographic details were published, the general consensus was that the active site was made up of a cubane-linked mononuclear nickel species. Therefore, mononuclear Ni complexes have been investigated as functional models for individual steps of the reactions mediated by the CODH/ACS [110]. Holm et al. have described an early example of a functional model of the ACS activity, using a tripodal NS3 ligand to form the Ni complex [NiCl(NS3)], 39 [111]. This work enabled the first characterization of a Ni-CH3 species 40 without strongly donating and stabilizing phosphine ligands, as well as the corresponding acetyl species 41. Coupling of the acetyl group with a thiol yields the desired acetylthioester. Met. Ions Life Sci. 2, 181–240 (2007)


205

Although this Ni species operates in a strict stoichiometric fashion – inactive metallic nickel is formed as a side-product – this system has provided useful insight into the putative reaction sequence of the ACS active site. Later work from the same group employed bipyridine instead of the biomimetic NS3 ligand. With this system, stable nickel complexes could be obtained, also after dissociation of thioester [112]. +

+

N MeMgCl

S Ni

R S R

Cl

+

N

N S

S R

Ni

R

S

S R

S

CO

Ni

R

CH3

R

R O

40

39

S

S

R

CH3 41 RSH

O

N +

S

RS

S

R S

CH3

+ Ni0

R

R

Riordan et al. more recently reported on a nickel complex 42 with a tripodal tris-thioether-borate ligand. Reduction of the easily accessible NiII complex with MeLi under CO atmosphere furnished the corresponding tetrahedral NiI-CO species 43, which showed a rhombic ESR signal with significant axial character and an IR-active νCO vibration at 1999 cm1 [113–115]. Although this latter value was consistent with the available data for the natural system, the differences in the ESR spectra indicated that the geometry around the reduced NipI center was not likely to be tetrahedral, but presumably trigonal bipyramidal or tetragonally distorted octahedral. O C

Cl S

Ni S

S

MeLi CO

S

Ni S

B

B

42

43

S

Most modeling efforts to date have focused on the dinuclear Ni center, disregarding the Fe4S4 subunit, which despite structural simplicity is known to induce significant difficulties in coordination chemistry attempts, especially if linkage is Met. Ions Life Sci. 2, 181–240 (2007)

206


sought through only one thiolate group [116]. Holm et al. have recently described some promising results that were aimed at linking a Fe4S4 cluster to a nickel center through only one sulfur bridge [117]. However, some progress in the preparation of Fe3S4Ni as well as Fe 4S4Ni complexes which could serve as models for the A-cluster was made, unknowingly, in efforts to actually model the ‘supposed’ C-cluster, prior to the elucidation of the unique structure of this C-cluster by X-ray crystallography. For instance, Pohl and coworkers [118–120] prepared Fe 4S4 cubanes linked to a square planar Ni center via either one or two thiolate bridgehead units, by changing the ligands on the Fe 4S4 cluster reagent. These species were prepared by reaction of a Ni{N2S2} metalloligand with [Fe4S4I4] 2 yielding 44, where the Ni center is coordinated to the Fe/S cluster in a bidentate fashion. Subsequent addition of KStip (Stip 2,4,6-tris(isopropyl)thiophenolate) gave the corresponding complex [(Ni{N2S2}2Fe 4S4 (Stip) 2] (45), in which each Ni is bridged through only one thiolate to the FeS cluster (monodentate), with a Ni–Fe distance of 2.827(1) Å. This complex exhibits a quasi-reversible oxidation wave at El/2 0.15 V (compared with SCE) [118–120]. X X

S

X

X

S

Ni Fe

S

Fe Fe S Fe I I S

S S

X + 2 KStip

Ni S

S Fe S S

X

X

S

Ni

S

S

Fe Fe S Fe S S

Ni S

X

X = NEt

44 45

The dinuclear bis(thiolato)-bridged dinickel fragment of ACS has been the subject of intense research in recent years, in part because of the scientific challenge to link two nickel centers with very different geometries via two mere thiolate units. Due to the initially unknown identity of the central metal atom – Cu, Zn, and Ni were considered – a few Ni-Cu model complexes have also been described. Despite the recent finding that the native enzyme only shows activity with Ni as the metal in the proximal site, while Cu and Zn inhibit catalytic activity, these complexes still have their respective merits, especially from a coordination chemistry perspective. The exact geometry of the so-called proximal metal (nickel) center, i.e., linked to the Fe4S4 cluster through a cysteine-bound thiolate group, remains unclear, with both square planar and tetrahedral coordination assumed plausible. Riordan et al. reported on the preparation and isolation of heterodinuclear thiolate-bridged NiIICuI complexes [121]. Prior to this report, only one example Met. Ions Life Sci. 2, 181–240 (2007)


207

of a thiolate-bridged NiCu dimeric species was characterized [122]. Simple addition of a cationic CuI salt such as [Cu(CH3CN) 4](BF4) to a diamino-dithiolate Ni metalloligand led to selective formation of a paddlewheel structure, while for the diamido-dithiolate Ni metalloligand 46 a tetranuclear complex 47 with a steplike structure was obtained. Notably, the proposed chelation of the Ni{N2S2} metalloligand was not observed, as two adjacent thiolate groups are coordinated to two different CuI ions, which is in stark contrast with results obtained with CuII-Ni{N2S2} bimetallic complexes. 2–

2–

O

O

O S

N

[Cu(CH3CN)4]BF4

Ni N

N

S

Cu

S

S

Cu

S

Ni

S

N

O

N Ni N O

O 47

46

Employing the bulky, tripodal borate-ligand PhTttBu (PhTttBu phenyltris((tertbutylthio)methyl)borate instead led to clean formation of the desired NiCu dimer 48 [112]. In this complex, the copper center adopts a distorted tetrahedral geometry, wherein one of the four Cu–S bonds is elongated, similar to the asymmetric coordination observed for the central atom in the native system. As the active site of the natural enzyme exhibits reactivity towards CO, which is apparent from ESR line broadening upon isotopic perturbation with 57Fe, 61Ni or 13CO as well as from an IR vibrational band for νCO at 1995 cm1, the reactivity of 48 towards CO was tested, but no features for a terminal CO bound to this dinuclear species could be observed. Rather, decomposition of the product with formation of mononuclear Cu(CO)(PhTttBu) occurred, together with reformation of the Ni{N2S2} metalloligand fragment [121]. 2–

O

2–

O S

N

Cu(CH3CN)(PhTttBu)

N

N

S Cu

Ni S

O

S

N

Ni

S

S B

S

O 46

48

Although many dithiolate-diamide ligands have been prepared, few examples of ligands with either unsymmetric or functionalized backbones have been synthesized and/or applied in coordination chemistry. Rauchfuss et al. developed Met. Ions Life Sci. 2, 181–240 (2007)

208


a strategy starting from the hexamethylated Ni{N2S2} metalloligand 49 [123], which showed similar coordination behaviour towards CuI salts as the system described by Riordan et al. [121]. 2+

S

N

[Cu(CH3CN)4]BF4

N

Cu

S

N

Ni

N

N

Cu

S

49

S

S

N

PiPr3

Ni

Ni S

S

N

N

NCMe Cu

Ni

PiPr3

S

51

50

A significant observation was that addition of phosphine to the tetranuclear NiCuCuNi ‘step’-complex 50 led to clean formation of the corresponding dinuclear complex NiCu(NCMe)(PiPr3) (51), with the Cu center in a tetrahedral geometry [123]. The labile NCMe ligand can be exchanged with MeNC but exposure to CO does not lead to any new products. A new bio-inspired unsymmetrical ligand, obtained by reaction of 3-mercaptopropionic acid and ethylenediamine, was also converted into the corresponding Ni-containing metalloligand 52 and similar CuI-based chemistry was carried out. A notable difference for the reaction involving phosphine was that no MeCN ligand was present, leading to a trigonal planar NiIICuI-PiPr3 complex 53, which showed no reactivity towards CO. The increased donor character of the {N2S2} ligand might be responsible for the observed alteration in the coordination mode of the CuI ion [123].

2–

O S

N Ni N

–

O

S

Cu PiPr3

Ni N

PiPr3

O

S

N

[Cu(CH3CN)4]PF6

S

O 52

53 Ni(cod)2 CO (1 bar)

O S

N

Ni

Ni N

S

O 54

Met. Ions Life Sci. 2, 181–240 (2007)

CO CO


Figure 10.

209

Molecular structure of 54 [112].

Reaction of the Ni{N2S2} metalloligand 52 towards Ni(cod) 2 under a CO atmosphere yielded the spectacular dinuclear Ni2 species 54 (Figure 10), which encompasses both a square planar NiII and a tetrahedral Ni0 center as well as terminal CO ligands bound to Ni, with IR bands for νCO at 1948 at 1866 cm1 [123]. X-ray crystallographic analysis revealed a Ni–Ni distance of 2.8 Å and slightly longer Ni–S bond lengths for the tetrahedral Ni0 (⬃2.35 Å) compared with square planar NiII (⬃2.2 Å). In the natural system, however, only one terminal CO ligand is bound to the Nip site, and the corresponding IR band comes at higher wavenumber. This might well imply a different oxidation state of this Ni center, as Ni0 is unlikely to play a role in any biological system. Complex 54 is unstable upon addition of donor ligands or introduction of air, leading to degradation of the complex and formation of well-defined mononuclear nickel species [123]. A faithful model concerning the reproduction of a peptide backbone, as in the native system, was developed by Krishnan and Riordan very recently [124]. They realized that cysteine functionalities in synthetic peptide-based ligands for nickel complexes were rarely utilized, although these electron-rich species are wellsuited for the stabilization of higher oxidation states of Ni (i.e., Ni III). Therefore, a Cys-Gly-Cys tripeptide motif 55 was devised for the synthesis of the Ni{N2S2} metalloligand 56, which was further metallated to yield a dinuclear model for the ACS active site [124]. The transformation of the apo-peptide to the desired Ni complex was established by its UV/Vis spectrum, which showed similar features as for reported diamido-dithiolate complexes, as well as by NMR spectroscopy and ESI-MS analysis. Addition of NiCl2·6H2O to this metalloligand or direct addition of Ni(OAc)2 and base to the apo-peptide ligand yielded a trinuclear species [Ni(CGC)Ni(CGC)Ni] 2 (CGC the Cys-Gly-Cys ligand scaffold). However, upon reaction of the NiN2S2 metalloligand with NiCl2 (depe) (depe 1,2bis(diethylphosphino)ethane), the dimeric complex 57 was selectively obtained, as was shown by various spectroscopic methods.

Met. Ions Life Sci. 2, 181–240 (2007)

210

VAN DER VLUGT and MEYER NH2

NH2

O O

NH2

2–

O NH NH

SH SH

O

Ni(acac)2

O

O S

N Ni N

KOH

55

Ni

Ni N

P

S

N

S

O NHAc

O NiCl2(depe)

S

P

O NHAc 56

NHAc 57

Despite the absence of crystallographic data, both Ni centers in 57 are proposedly in a square planar environment. This NiIINiII complex undergoes two one-electron reductive processes, yielding a mixed-valence NiIINiI and a NiIINi0 complex, with the phosphine-coordinated nickel being reduced selectively. The parent complex 57 does not bind CO, but both reduced states do show pronounced interaction, as observed by cyclic voltammetry. Although the oxidation states for both Ni centers in the reduced form of the A-cluster are under debate, and a NiIINi0 mixed valence situation cannot be completely ruled out, the alternative NiIINiI [Fe4S4] seems more attractive [125,126]. Schröder et al. synthesized analogues of such square planar dinickel species, also employing chelating phosphines for stabilization and capping of the ‘Nip’-type site [127]. This proposed Nip-type site showed a distorted square planar geometry, with P–Ni–S angles of ⬃97. Furthermore, this nickel center could be reduced in a one-electron process at E1/2 0.47 V, to yield an ESR-active monocationic species. Also the groups of Holm [128] and Darensbourg [129] have reported on complexes with related Ni{N2S2} metalloligands and their binding affinities for Cu, Zn and Ni. Similar results to those published by Riordan have also been obtained by Mascharak et al., who employed the more rigid metalloligands 58 and 61 to obtain the Ni-dimeric complexes 59 and 62, respectively [130]. Upon reduction of the dimeric NiIINiII species 59 under CO, an ESR signal was observed for the NiI center, which is proposed to originate from the terpyridine-coordinated Ni site (60). In the corresponding IR spectrum, a vibrational band for νCO was discernible at 2044 cm1, typical for a terminal CO bound to a six-coordinate NiI ion. For complex 62, the analogous NiIINiI-CO species (63) featured an IR band at νCO 1997 cm1, which is in good agreement with the value observed for the CO-bound reduced form of the native enzyme at 1996 cm1. The so-called Nip center – the Ni center that undergoes reduction – also shows labile coordination, as excess phenanthroline results in formation of [Ni(phen)3] 2. Reaction of [CH3Co(dmgBF2)2L] (dmgBF2 (difluoroboryl)dimethylglyoximato, L py, PEt3) with two equivalents of [Ni(tmc)] gave [Co(dmgBF2)2L] , [Ni(tmc)] 2, and [Ni(tmc)CH3] in 80% yield [57]. The overall transformation Met. Ions Life Sci. 2, 181–240 (2007)


O

O

O N

S

N

N

S

S

N

N

N Ni

S

N N

O

N

59

58

S Ni

O

O

N

N

Ni

Ni

Ni

CO

Na2S2O4 CO (1 bar)

N

S

N

NiCl2(terpy)

211

60 N = 2,2',6',2"-terpyridine

N

O

O N

S

O N

NiCl2(dppe)

N

N

S

61

S

P

S

P

N

P

Ni

Ni

O

O

CO N

Ni

Ni

Ni

P

S

Na2S2O4 CO (1 bar)

S

P

O

62

63

= dppe = 1,2-bis(diphenylphosphino)ethane P

provides a model for the transfer of a CH3 group from methylcobalamin to the nickel ion of CODH. R,R,S,S-[Ni(tmc)CH3](BAr4) (BAr4 B(3,5-(CF3)2C6H3)4 ) was characterized by X-ray diffraction [57] (Figure 1).

3.3. Nickel Superoxide Dismutase The latest addition to the growing list of nickel-containing metalloenzymes with S-coordination is nickel superoxide dismutase (NiSOD) [131], for which very recent crystallographic analyses of samples obtained from Streptomyces species and various cyanobacteria unequivocally determined the presence of Ni, thereby establishing the identity of a fourth subclass of SOD enzymes, after the Fe, Mn, and Cu/Zn SODs were already known. The active site has been characterized in both a reduced and an oxidized form, with major structural differences existing between these two states [132,133]. One feature that sets this active site apart from the other SOD metalloenzymes is the ligation of thiolates, and it remains unclear how sulfur-based oxidation is suppressed during the catalytic turnover of O2•. For a full account of the results of studies on the native active site, see Chapter 10 of this volume. Met. Ions Life Sci. 2, 181–240 (2007)

212

VAN DER VLUGT and MEYER H N

HN N O N H2N

NiII

S

SCys reduced

O2 – 2 H+

H2O2

O2

O2 –

N O N H2N

NiIII S

SCys oxidized

Superoxide is proposed to bind at the sixth open coordination site, trans to the histidine binding site. The significant role of the histidine ligand in the catalysis was examined by site-specific mutagenesis, which showed dramatically decreased activity upon alteration at this position. The square planar NiII site has been adequately modeled (unknowingly) by several groups, in efforts to mimic the distal Ni-site in ACS. One of the earliest examples was provided by Krüger, Peng, and Holm, who described diamido-dithiolate nickel complexes as models for the [NiFe]-hydrogenase active site [134]. Despite the obvious similarities with the aforementioned Ni-containing active sites, the presence and active participation of a histidine fragment during the catalytic conversion of superoxide into H2O2 (oxidation of NiII to NiIII) or O2 (reduction of NiIII to NiII) has so far eluded accurate structural or functional models. More than ten years before the elucidation of the NiSOD structure and the realization that Ni-containing enzymes can convert O2•, two Japanese groups collaboratively reported on the use of NiII complexes with oligopeptide ligands for the ‘dismutation’ of both O2• and H2O2 [135]. First, a characteristic ESR signal was generated by addition of the spin-trapping agent DMPO (DMPO 5,5-dimethyl-1pyrrolyl N-oxide) to a solution of in-situ generated O2• ions. This signal completely disappeared upon addition of native SOD enzyme (the precise nature of this metalloenzyme was not specified). Out of eight nickel(II) complexes with coordinated tri- and tetrapeptides, those containing a histidine unit in the third position of the amino acid sequence showed surprisingly high activity upon introduction to identical O2• solutions, as indicated by large reductions of the ESR signal for the DMPOO2• system. Upon addition of H2O2 to solutions containing the same NiII -peptide complexes with His in the third position of the amino acid sequence, the DMPOO2• ESR signal was detected. The presence of O2• ions was also supported by UV/ Vis spectroscopic monitoring of the formation of a blue pigment called formazan, which results from the oxidation of NBT (para-nitro blue tetrazolium chloride). Few well-defined model complexes with an NiIII{N2S2}(N) coordination are known, and the preparation of stable NiIII species is more difficult than for isostructural CuIII complexes, as the NiIII /NiII redox potentials are more positive (see also Chapter 3). Furthermore, decomposition of most NiIII systems is rapid. For NiIII thiolate complexes, autoreduction of the nickel ion by the thiolate ligands, concomitant with formation of disulfide species, is often encountered. Met. Ions Life Sci. 2, 181–240 (2007)


213

Several groups have described the generation of such NiIII species with an {S4} or an {N2S2} ligand set by electrochemical methods, without successful isolation of the corresponding complexes as solids [136,134]. The most elegant species was reported by Hanss and Krüger in 1998 [137]. The four-coordinate NiII complex (64) was formed by addition of Ni(OAc)2·4H2O to the ligand (H4phmi). 2–

O

O NH NH

SAc

Ni(OAc)2.4H2O

N

OH–

N

SAc

S Ni S

O

O (H4phmi)

64 electrochemical oxidation –

O py S

N

–

O pyridine

N

Ni S

O

S

N

Ni N

S

O 66

65

This red NiII complex with square planar geometry was studied by cyclic voltammetry, and a reversible one-electron oxidation occurred at 0.71 V (compared with Fc/Fc). Subsequent quantitative bulk-electrolysis yielded compound 65, for which the corresponding molecular structure showed little structural changes compared to the NiII species 64 – the intramolecular S–S distance decreased from 3.224 to 3.069 Å, but no direct bonding interaction was observed – indicating that the oxidation is mainly nickel-centered. Complex 65 exhibited a magnetic moment of 1.78 µB, which relates to a low-spin NiIII center. In agreement with this observation, no ESR signal was detected at 100 K. However, upon addition of excess pyridine to a solution of 65, the ESR spectrum of complex 66 showed a nearly axial signal with g values at 2.313, 2.281, and 2.000. Furthermore, three additional lines of equal intensities (ANz 25 G) were observed, due to hyperfine coupling with the axially coordinated nitrogen ligand pyridine. Although this system was not intended at the time as a faithful structural model for NiSOD, the observed ESR spectrum shows great similarity to the spectroscopic data obtained for the native NiSOD system [133] (Figure 11). An elegant approach to combine structural modeling with the development of a functional model was disclosed very recently by Shearer and Long [138]. Based Met. Ions Life Sci. 2, 181–240 (2007)

214


Figure 11. ESR spectra for (A) complex 66 at 100 K in DMF [137] and (B) for oxidized NiSOD from S. coelicolor at 55 K [133]. Reproduced with permission. Part A: Copyright (1998) Wiley-VCH, Weinheim. Part B: Copyright (2004) American Chemical Society.

on the known peptide sequence for the active site of the native enzyme, solid phase peptide synthesis on Wang resin was employed to prepare a 12-residue peptide chain reminiscent of the N-terminus of the active form of S. coelicolor NiSOD. This apo-peptide was then coordinated to NiII in an equimolar ratio by reaction with NiCl2 at a pH of 7.2 (at pH 6, no binding of Ni was observed). No structural data were provided for this system, but UV/Vis and CD spectra resembled those recorded for the natural system. Oxidation of the NiII center does not occur, even when kept in air, as indicated by the absence of any ESR signal. However, catalytic conversion of O2• was observed after addition of solid KO2 to a buffered solution of the metal complex: effervescence indicated the formation of O2, while H2O2 could be assayed with an appropriate kit. For every mole of O2• added, ⬃0.5 mol H2O2 was produced. Further evidence for SOD activity was provided by the observation that the oxidation of NBT by O2• was effectively suppressed in the presence of the Ni complex, up to at least ten thousand equivalents of O2•.

4.

MODELS FOR THE UREASE ACTIVE SITE

Urease is a unique metallohydrolase that contains two proximate NiII ions within its active site (see Chapter 6 of this volume). It efficiently catalyzes the hydrolysis of urea to give carbon dioxide and two moles of ammonia as final products. After the first X-ray crystal structure of microbial urease was published in 1995 [139], accumulating crystallographic evidence has meanwhile provided a detailed picture of the Met. Ions Life Sci. 2, 181–240 (2007)


215

active site of the enzyme and a solid background for an understanding of its mechanism of catalysis [140,141,142]. The initially proposed mechanism of urease activity assumed that urea is first activated by coordination to one NiII ion, in conjunction with extensive hydrogen bonding within the active site pocket of the protein, and is subsequently attacked by a nucleophilic hydroxide terminally bound to the opposite NiII ion [143,144]. The resulting carbamate would then decompose further to produce carbonic acid and another molecule of ammonia (Scheme 6, left sequence). NLys O

O

HisN HisN

H2N

NHis

O H

OH2

O

NHis Ni

Ni

Asp

O

H2O

O

NH2

O H2N

NH2

2 H2O 2 H2O

NLys O HisN HisN

NLys NHis

O

Asp

O

O H

O

O

NHis

Ni

Ni

HisN

Ni

H2N

NHis NHis

Ni

H O

Asp

O

O

O

NH2

O

HisN

NH2

O

NH2

NH3

NLys O HisN HisN

NLys NHis

O

O

O NH2

O Asp

O

O

HisN

NHis

Ni

Ni

HisN

Ni

H O O

O

NHis NHis

Ni

Asp

O NH2

O

NH2

CO2 + NH3

CO2 + 2 NH3

Scheme 6. Proposed mechanisms of urease action. Met. Ions Life Sci. 2, 181–240 (2007)

216


Based on the crystal structures of urease inhibited by either phosphate, diamidophosphate or borate [142,145], as well as on the basis of recent model calculations [146,147], an alternative mechanism has been proposed and the sequence of events outlined in the right part of Scheme 6 now seems most likely. It implies that urea first coordinates to both nickel ions in a N,O bridging mode, followed by nucleophilic attack by the bridging hydroxide. However, details of this process, including the mode of urea binding and the exact identity of the nucleophile, are still under debate, and cyanate has also been proposed as a possible intermediate in the urease mechanism [148]. The very slow, uncatalyzed decomposition of urea gives ammonia and cyanic acid with a half life of 3.6 yr at 38C in the pH range 2–12 [149–151]. As a consequence, the design and investigation of dinuclear NiII model complexes that mimic characteristics of the urease active site and that are capable of binding and degrading urea receives particular current attention [152]. Some specific active site structural features of urease are the asymmetric N/O-rich coordination environment, with a Ni–Ni distance of around 3.5 Å, the bridging carbamate (often modeled by a bridging carboxylate), and the presence of a hydrolytically active Ni-bound hydroxide or water. The ability to bind urea is a prerequisite for urease-like activity, and different urea binding modes have been observed in synthetic model compounds. Complex 67 [153] containing a ( µ-aqua)bis( µ-carboxylato) core reacts with urea to provide the fi rst example of urea coordination to a dinickel(II) site [154]. In the resulting species 68, the urea molecule binds through its carbonylO to one of the Ni II ions, which is the most commonly observed coordination mode.

Me2 N

O

H

O O

O N Me2 Ni O Ni Me2 N H O N O O Me2 O

+ O H2N

NH2

Me2 N N Me2

O

O O

Ni O O

O Me 2 N Ni N O Me2

H2N 67

OTf –

NH2

68

More recent studies have employed well-designed dinucleating ligand scaffolds for proper orientation of the two proximate NiII ions. Compartmental ligands based on bridging alkoxide [155–159], phenolate [160–164], pyrazolate [165–171] or phtalazine units [172–174] are the most prominent ones. Selected examples are 69 [160] and 70 [158], in which the two nickel ions are additionally spanned by carboxylate bridges. Magnetic coupling between the two high-spin NiII ions is usually anti-ferromagnetic in those cases. Met. Ions Life Sci. 2, 181–240 (2007)


217

+

2+

Et

Et

N

N

Ni

N N

N

N

Ni

O O

Et

Ni

N

N

N N

O O

N

O N

N

O

N

N

N

N

O

O

Et

N

OH2

H2O

N

N

Ni

70

69

Some emphasis has been placed on emulating the asymmetry of the donor environment of the urease active site in synthetic model complexes [163,169,175]. Complexes 71 [176] and 72 [159] are examples that feature solvent-filled accessible coordination sites. Some of these asymmetric phenolate- or alkoxide-derived dinickel systems were shown to promote the hydrolytic cleavage of the activated substrate 4-nitrophenylphosphate [156,159,163], but none has been reported to exhibit hydrolytic activity towards urea or amide model substrates. +

N Ni

N N

N

O

O

N

O

Ni

Ni

N

O O O O

OH2

N

Ni

N

N O OH Et

HO O Et

71

+

OClO3

N

72

A bridging hydroxide is present in 73 [177], 74 [167], and 75 [165], and complex 73 has been tested in the catalysis of bis(p-nitrophenyl)phosphate hydrolysis. A plot of the rate constant against pH showed a sigmoid curve with an inflection point at a pH close to the pKA of the Ni-coordinated water (⬃8.5), which is consistent with the metal-bound hydroxide acting as the nucleophile [177]. 2+

Ni N

Me2 N

N

N N

Me2 Me2 N H N

O O H

73

N

Ni N

N

Ni

N Me2

2+

Me2 N N N Ni O H

74

3+

N N Me2

N N

N

N

Ni

Ni

N O Me2 N H C Me

N C Me

N Me2

75

Met. Ions Life Sci. 2, 181–240 (2007)

218


Complex 75 is of some interest since it provides additional N-functional groups in proximity to the bimetallic core. During complex formation, these basic amines pick up a proton to generate the Ni-bound hydroxide at neutral pH, which mimics essential characteristics of the protein surrounding in the urease active site [165]. At the same time, accessible sites are available for substrate binding, as the acetonitrile ligands in 75 are easily replaced by DMF or N-bound thiocyanate [165]. However, no hydrolytic activity towards urea could be observed. The bridging H3O2 moiety in 76 is hydrolytically more potent than the tightly bound bridging hydroxide in the related complex 74 [167]. In contrast to the latter, 76 is capable of nitrile hydration to give complexes 78 with N,O-bridging amidato groups which are assumed to be generated via initial replacement of water by the N-bound substrate within the bimetallic pocket (77). This picture is corroborated by the molecular structure of 79, which closely mimics the situation in the putative intermediate 77 [178].

N

2+

2+

NEt2

NEt2

Et2N N N

Ni

RCN

N Ni

O H O N H Et2 H

∆

N

Et2N N N

Ni

N N Et2 C

N Et2

N

Ni O H

N Et2

R 76

77

2+

NEt2

Et2N

N Ni N N Ni N N Et2 C

NEt2

N

O

N Et2

Et2N N N

N Ni

Ni

N HN Et2

O

N N Et2

R 79

78

The phtalazine-based system 80, reported by Barrios and Lippard [172], features bridging water molecules that exhibit rather low pKa values of 6.49 and 9.71. While 80 itself is hydrolytically inactive, the singly deprotonated form 81 mediates the hydrolytic cleavage of picolinamide (but not acetamide) in ethanol to finally give 83 [172]. Kinetic data and IR spectroscopy suggest a substrate binding pre-equilibrium with a binding constant of picolinamide to 81 of 70 ± 20 M1, giving complex 82. Subsequent nucleophilic attack is assumed to occur by the bridging hydroxide, which appears to be suitably arranged when the picolinamide-O is located in a bridging position. Met. Ions Life Sci. 2, 181–240 (2007)


219

4+

N N

N Ni

N

O H2

H2O

NaOH

N

H2 O Ni

3+

N N

N N

N OH2

H O

Ni

N Ni

O H2

H2O

N OH2

81

80 O

O NH2

EtOH

EtOH

NH2

2+

N N

N Ni

N N

N Ni

O O O O

83

N N

2+

N N

N Ni

N N

H Ni O O

N N OEt H

NH2 82

Molecular details of urea molecules attached to dinickel(II) cores have now been elucidated for several systems. After the initial report of 68, η1-O bound urea has been crystallographically confirmed in the (µ-alkoxide)(µ-carboxylato) complex 84 described by Yamaguchi and coworkers [157], in a few phenolate-bridged species reported by Okawa and coworkers (such as 85 [161] with a mixed-spin state) and by Nordlander’s group (Ni4 complex derived from unsymmetric ligands [163]), in several pyrazolate-based dinickel(II) compounds with additional carboxylate bridges and either symmetric (86, 87) or unsymmetric (88) cores (for the molecular structure of 88 see Figure 12) [169], and in the phtalazine-derived complex 89 [173]. In the latter two cases, species with N-substituted urea have also been structurally characterized [169,174], as has a phtalazine-based dinickel complex with S-bound thiourea [174]. Whereas 87 provides additional terminal sites for solvent coordination [169], 89 incorporates both a µ-aqua and a µ-hydroxide group [174]. In 86, 87, and 88, urea coordination is stabilized by H bonding between the urea NH fragments and the O atoms of the bridging acetate [165], supporting the ideas of crucial H bonding patterns in the enzyme active site. Type 89 complexes feature Met. Ions Life Sci. 2, 181–240 (2007)

220


urea H bonding interactions with the bridging hydroxide [173,174]. All NiII ions with η1-O bound urea are usually six-coordinate. 2+

H2N N N

N

O

Ni

RHN

OH2

N Me2

85 2+

Et S N N

Ni

Ni

S O O Et H N H

N

Ni

O H

N Me2

N

NH2 84 Et S

N

O Ni

O

OO H2N

N

Ni

N

NH2

O

N

H MeO

O S O Et H N H

N N

N

N

N Me2 NHR

Ni

Ni O

O

O

O

H N Me

HN Me

86

2+

H OMe N N Me2

H N Me

NH Me

87 3+

Et2 N

Et S N

RHN

N N Ni

Ni

S O O Et N H

O

2+

N

N N

N N

N Et2

H H2N 88

N

Ni H Ni N O O O H2 O NH2 H2N

NH2

89

Two examples 90 and 91 are known where urea forms a single atom bridge between the proximate metal ions through its carbonyl-O atom [173,179] (Figure 13). An N,O-bridging coordination of (deprotonated) urea occurs in 92 [166], which forms in an equilibrium reaction between the active H3O2-bridged species 76 and the substrate (K1 4.3 ± 0.4 M in acetone, 2.7 ± 0.5 M in MeCN) [171]. Kinetic studies by stopped-flow methods provided insight in the process of urea incorporation into the bridging position: binding was shown to follow a rate Met. Ions Life Sci. 2, 181–240 (2007)


221

Figure 12. Molecular structure of the cation in complex 88 [169].

nonlinear in urea concentration, indicating that two urea molecules are involved. A detailed kinetic scheme for the multi-step process of urea binding to 76 has been derived. Steady-state intermediates of the tentative formula [LNi2 (OH)(urea) n] 2 are formed that are close analogs of the reactive intermediate of urease, which also has the hydroxide and urea ligands bound at the bimetallic array [140,144]. However, the hydroxide then acts as a base in the model system, whereas it attacks as a nucleophile in the enzyme. Starting from the related H3O2-bridged dinickel

Figure 13. Molecular structure of the cation portion of complex 90 [173]. Met. Ions Life Sci. 2, 181–240 (2007)

222


complex 93 with triazacyclononane side arms attached to the pyrazolate, not only parent urea (see molecular structure in Figure 14) but also N-substituted urea derivatives could be incorporated in the N,O-bridging position [170]. 3+

Bu

N N

N Ni

N

H O

Ni

Bu

N C Me

O

H2N

N

O

O

Bu

O NH2

NH2 91 (L = MeOH)

90 2+

NEt2

Ni

H2N

NH2

Bu

N

O

O

O

L

L Ni

N

O H2N

N

N

N N Ni

H2N

N

Ni

2+

NEt2

O

Et2N

NH2

N

Et2N

N N Ni

Ni

N

H2O

N O H O Et2 H H

N HN Et2

N Et2

O

N Et2

NH2 92

76

An unprecedented µ3-κN:κN:κO triply bridging coordination mode of deprotonated urea has been observed in the tetranuclear NiII complex 94, which assembles from two pyrazolate-based bimetallic entities [168]. In 94, the presence of two terminal four-coordinate low-spin NiII and two central six-coordinate high-spin NiII leads to a mixed-spin situation, while results from UV/Visspectroscopy and magnetic measurements in solution suggest the coordination of additional solvent ligands to the outer metal ions at low temperature [168]. H2N

Me N N N H H Ni N N O NMe2 H N Me2 Me2 N O Me H 2N N N Ni H H Ni N N N O NMe Me Me

iPr

iPr

N

N

N N

N

Ni N

iPr

2+

N

Ni

O H

H O H

N

iPr

93

Met. Ions Life Sci. 2, 181–240 (2007)

4+

NH2

H2N

N O Ni

NH2

94


223

Figure 14. Molecular structure of triazacyclononane-derivative of structure 92. This compound was formed by reaction of H3O2-bridged species 93 with urea [170].

It is generally believed that the hydrolytic stability of the urea molecule is due to its high resonance stabilization energy (estimated to be 30–40 kcal/mol), which presumably is reduced upon coordination to metal ions, activating it for nucleophilic attack. In order to assess the activation of the urea substrate in its different binding situations at dinickel sites, changes of the CO and CN bond lengths as well as in the ν(CO) stretching frequency upon coordination are usually considered diagnostic for the weakening of those bonds or polarization of the carbonyl moiety. A comparison of the structural data available for dinickel(II) complexes with bound urea (or substituted urea) is illustrative, as summarized below in Scheme 7. Coordination of urea to a single NiII via its O atom (mode A) does not appreciably alter the metric parameters of the carbonyl group and causes only a moderate shift of νCO from 1683 cm1 [180] to around 1640–1660 cm1, suggesting that the substrate is poorly activated towards hydrolysis in this binding mode. However, the two C–N bonds may become significantly different in the bound substrate, which is particularly obvious for the only complex to date, 84, that shows an appreciable catalytic activity in the ethanolysis of urea (see below). One may assume that urea would be better activated by O coordination to two NiII ions (single-atom bridging mode B), but data for the two known complexes that exhibit such a bonding mode only partly support this view. While the CO bond is somewhat lengthened in B, the C–N bonds are even shorter than in free urea. Few examples are known for urea spanning two NiII ions, though the substrate is deprotonated in those cases (mode C; some related diiron(III) complexes have also been reported). Such µ1,3-N,O bidentate coordination C indeed leads to partial loss of resonance as witnessed by a significant elongation of the distal C-NH2 Met. Ions Life Sci. 2, 181–240 (2007)

224


NH2

H2N

Ni

Ni

O 1.258(1) Å

1.341(1) Å

H2N

O 1.24 - 1.27 Å

1.30 - 1.36 Å

NH2

1.31 Å HN

(A)

Ni Ni

Ni O

1.29 - 1.31 Å NH2

1.30 - 1.34 Å (B)

NH2

O

NH

1.43 Å H2 N

O Ni

(D)

O

HN

1.26/1.29 Å H2N

Ni

Ni

Ni

Ni

1.25 Å

~1.27 Å

NH2 1.35 - 1.39 Å

(C)

Scheme 7. Comparison of structural data for dinickel(II) species A–D, containing coordinated (substituted) urea.

bond. However, the anionic nature of the ureate should render it more resistant to hydrolysis, and accordingly no hydrolytic cleavage but release of ammonia and formation of cyanate was observed upon heating these compounds (see below). It certainly would be most favorable if urea coordinates to the bimetallic core in the µ1,3-N,O mode without deprotonation, just like in a CoII complex that revealed a more pronounced elongation of the C-NH2 bond to 1.42 Å [181]. This value is similar to that found for a unique µ3-N,N,O ureate in 94 (mode D), which may point towards the importance of secondary interactions such as hydrogen bonding with protein side chain residues (instead of interaction with a third NiII in 94) that contribute to the polarization of the substrate molecule. These interactions may also modulate the pKa of the urea upon coordination to avoid its premature deprotonation: while O-coordinated urea has almost the same pKa as the free molecule (around 14) [182,183], N coordination has been shown to dramatically increase urea acidity (down to pKa 3 in aqueous solution) [184], and this should be even more pronounced in the µ1,3-N,O mode. Even if one considers the relatively lower polarizing power of NiII compared with the metals used in those studies, deprotonation is still likely to occur upon bidentate coordination under conditions optimal for urease (pH 4–8) if modulating interactions are not present (such as in the model systems currently available). It should be noted that N-coordination of nondeprotonated urea to 3d metal ions (including Ni2) has been achieved with chelating N-(2-pyridylmethyl)urea [185]. In the presence of NiCl2, this substrate undergoes ethanolysis or hydrolysis while zinc ions are ineffective [186,185]. Met. Ions Life Sci. 2, 181–240 (2007)


225

In view of these considerations it may be not surprising that none of the model complexes mediates the direct hydrolytic degradation of parent urea. Ethanolysis of urea in a slow catalytic reaction (4 equivalents of ethyl carbamate after 12 h at 80C) has been reported for 84 [157]. Interestingly, the related hydroxobridged dinickel complex is not active, but readily absorbs CO2 to give the complex bridged by hydrogencarbonate (HCO3 ). The latter again exhibits ethanolysis activity similar to the acetato-bridged species 84, which resembles the activation of apo-urease in the presence of NiII ions by CO2. Compound 87 produces 2.2 equivalents of ethyl carbamate after 6 days at 80C [169]. On the other hand, several complexes have been shown to promote the elimination of ammonia from coordinated urea to yield metal-bound cyanate. This has been taken as indication that a cyanate intermediate may also be relevant for the enzyme, and a high-level computational study by Estiu and Merz on urea decomposition promoted by Lippard’s phtalazine-based dinickel model complexes led the authors to propose that both the elimination and hydrolytic pathways may indeed compete in the urease active site [187]. The first experimental evidence for the transformation of urea to cyanate at dinickel(II) sites came from the groups of Okawa [162] and Meyer [167] for O-bound urea or N,O-bridging ureate, respectively. A series of dinuclear NiII complexes derived from an unsymmetric phenolate ligand (such as 95) react with urea in refluxing ethanol solution to give dinickel products 96 with N-bridging cyanate in low yields (17–23% after heating for 24 h) [162]. Heating of 92 gives rise to ammonia extrusion from

+

N

+

O

N

O

N H2N

Ni

N N

Ni

O O O O

NMe2

Ni

N

in EtOH reflux

N

OMe H

2+

OEt H

O

Ni

- NH3

N Et2

Et2N N N

N

N Ni

N HN Et2

O

2+

NEt2

Et2N

Ni

N C

NMe2

O 96

N N

N

Ni O

95 NEt2

N

O

NH2

N Et2

N Ni

N

C O

N Et2

H2N 92

97

Met. Ions Life Sci. 2, 181–240 (2007)

226

VAN DER VLUGT and MEYER O

Ni

Ni

+ H2N

NHR

– 2 H2O

Ni

Ni O

O H O H H 93

NH NHR

MeOH / H2O / ∆

(R = H, Ph, CH2Ph)

O + H2N

OMe ∆

– 2 H2 O

Ni

Ni O

– NH2R

+ NaNCO

MeOH / ∆

Ni

Ni

NH

O C N

OMe

+ KNCO

Ni N

Ni O C N

C O

Scheme 8. Transformations at the dinickel(II) complex 93 relevant to the urease active site [170].

the N,O-bridging ureate and formation of the cyanate-bridged complex 97 [167]. However, subsequent hydrolysis of the formed cyanate could not be observed with those systems, even when the more stable pyrazolate complex with triazacyclononane compartments was used that allows investigations in aqueous solution. Conversions at the bimetallic scaffold of 93 relevant to urease are summarized in Scheme 8 [170]. Both complexes 89 and 90 promote the hydrolysis of urea in a two-step process [173]. Heating of 89 or 90 in acetonitrile solution produced ammonia with kinetic first-order dependence on complex concentration and an observed rate constant of (7.7 ± 0.5) 104 h1 to yield a cyanate complex as the reaction product. It remains unclear, however, which binding mode of urea (terminal or bridging as found in 90) facilitates the elimination reaction. Ammonia elimination from the O-bound terminal substrate appears to be in accordance with quantum chemical studies on that model system [187]. When the reaction was carried out in 50% aqueous acetonitrile solution, ammonia was produced at the same rate but without buildup of the cyanate-containing product, suggesting that the latter is Met. Ions Life Sci. 2, 181–240 (2007)


N

N Ni

N

H O

NH3

N Ni

O H2

H2O

N

N

Ni

N

N

N H N 2 O

N Ni

O H2

H2O

O

227

N O C

H2N

NH2

H2O

N

H2O

N Ni

N H2O

N H N O O H2

N

N Ni

Ni

N

N

H2O

OH2

N H N 2 O

N Ni

O H2

N O

CO2 + NH3

O

NH2

Scheme 9. Proposed mechanism for the hydrolysis of urea with complexes 89 and 90.

hydrolyzed in the presence of water. Hydrolysis of the cyanate is also first-order in water (second-order rate constant of (9.5 ± 1) 104 M1 h1, unaffected by pH changes from 6.5 to 8.5), thus indicating that it proceeds by attack of an external water on the coordinated cyanate [173]. A mechanistic sequence as sketched in Scheme 9 has been proposed. In an analogous reaction, urea substrates having alkyl substituents at only one of the N atoms undergo alkylamine elimination to form a dinickel cyanate complex [174]. In contrast, no reaction was observed for N,N-dialkylated substrates. The results of most model studies for Ni-mediated urea degradation reported to date are consistent with a cyanate intermediate. While this differs from the most likely mechanism of urease activity as deduced from protein crystallography, there is still no definitive evidence ruling out a transient Ni-bound cyanate intermediate for the enzyme. Crystal structures of dinickel complexes 98 and 99 that incorporate O,O bridging or O,N bridging (O-methylated) carbamate have been discovered [166,170]. These are of interest because a carbamylated lysine residue spans the two nickel ions in the urease active site, and also because Ni-bound carbamate has been postulated as an intermediate in the original urease mechanism (compare Scheme 6). Complex 98 was found to produce one equivalent of ammonia upon heating in methanol/water solution [170]. A dinickel(II) complex of a tetrakis(benzimidazolyl)-substituted 2-hydroxy-1,3-diaminopropane ligand (similar to the ligand in 70) was reported to readily absorb CO2 in methanol solution to give a monomethyl carbonato-bridged species [188]. Met. Ions Life Sci. 2, 181–240 (2007)

228

VAN DER VLUGT and MEYER iPr

i

N

Pr N

N N

N

Ni N

Ni N

iPr

iPr

Pr N

N N

N

N

O

i

N

N

Ni O

iPr

iPr

2+

H2N

2+

N

Ni

NH i N Pr

O MeO 99

98

Acetohydroxamic acid (AHA) is known as a potent inhibitor of urease, and AHA coordination to dinickel sites has been emulated in 100 and 101, which are readily prepared from the reaction of 67 or 68 with AHA [189]. The inhibitor features a bridging binding mode with its hydroxamate-O symmetrically spanning both metal ions and the carbonyl-O chelating to one of the NiII. A similar situation is observed in 102 [190] as well as in complexes with salicylhydroxamic acid 103 [191], benzohydroxamic acid [192] and N-phenylacetohydroxamic acid [193]. This now appears to represent the general mode of hydroxamic acid coordination in metalloenzymes with dinuclear active sites and in their bimetallic model systems. In fact, 103 was reported prior to the crystal structure of AHAinhibited urease and led to the proposal of the hydroxamate inhibitor bridging the two nickel ions, as was later confirmed for the enzyme [194,195]. Compound 104 containing a deprotonated bridging N-hydroxyglutarimide was obtained in an unusual hydroxylamine elimination and cyclization reaction upon reaction of 67 with glutarodihydroxamic acid [196]. Four distinct hydroxamate binding modes have been observed in a heptanuclear NiII complex with +

Me2 O N Ni N Me2 O O N H

O Ni O

O Me2 O O Me2 O N N Ni Ni N O N Me2 O O Me2 NH NH2 H2N

OTf –

Me2 N N Me2 O

N H

N Ni

N

+

O Ni O

O NH

OTf –

O

O

O

OTf –

101

100

O

+

py py

O Ni

N N

102

Met. Ions Life Sci. 2, 181–240 (2007)

Ni O

O

O

O

O N H

N H O

+

py py O

N Me2

O

O

Me2 N

O Ni O O

N

HO

103

104

OMe2 N Ni N Me2 O

OTf –


229

2 (dimethylamino)phenylhydroxamic acid (2-dmAphaH), [Ni7(2-dmAphaH-1)2 (2-dmApha) 8 (H2O)2]SO4 ·15H2O [197].

5.

MODELS FOR ACIREDUCTONE REDUCTASE

Acireductone dioxygenase has recently been identified as a nickel-containing enzyme that catalyzes the oxidative breakdown of 1,2-dihydroxy-3-keto5-methylthiopentene (acireductone) in a CO-releasing process to give 2-methylthiopropionic acid and formic acid as part of the methionine salvage pathway [198] (compare Chapter 12 of this volume). From EXAFS and paramagnetic NMR data in conjunction with homology modeling, it was concluded that the resting state of the enzyme contains an octahedral NiII center ligated by three histidine-N and one glutamate-O, with two water molecules filling the remaining sites. In the presence of bidentate substrate the nickel is assumed to retain the octahedral environment with release of one histidine ligand [199, 200]. NHis H2O

NHis

Ni

NHis

H2O

OH

O

OGlu

S

S

OH

OH + CO +

H

OH

O

O

This chemistry has recently been emulated by Berreau and coworkers using a mononuclear NiII complex 105 of a bulky tris(2-pyridylmethyl)amine-type ligand and 2-hydroxy-1,3-diphenylpropan-1,3-dione as the test substrate [201]. Treatment of 105 with base gave a new (as yet unidentified) species; upon addition of O2 complex 106 is obtained in a reaction reminiscent of the ARD chemistry. When 18O2 was used, 106 was formed with a single 18O atom in each of the benzoate ligands, similar to the outcome of isotopic labeling studies with the native enzyme. Ph

HO

Ph

+

O

Ph Ph

O

H H O

CH3CN, O2 Me4NOH·5H2O

N

Ph O

O

Ni N

N N

O N Ni

O

CO

N N

Ph N

105

+

Ph

106

Ph

Met. Ions Life Sci. 2, 181–240 (2007)

230

6.


CONCLUDING REMARKS

Synthetic model studies for the different nickel sites in biology have not only uncovered exciting coordination chemistry, but have also provided significant insight into electronic structures and geometric requirements of individual species that have been proposed for the various mechanistic schemes. However, many details of the catalytic mode of action and the interplay of active site components such as the Fe4S4 cluster and the two nickel ions in ACS are still not fully understood. The availability of structural information at atomic resolution for the cofactors will certainly stimulate the design of even more sophisticated small molecule model systems to answer open questions, involving elegant synthetic approaches with either short metallopeptides or more classical small molecules. In addition, the search for new types of catalysts that mediate fundamental reactions such as hydrogen evolution based on the metalloenzyme functional principles is going to be a major task for the future. Just to mention urease as an example, it is amazing that a truly catalytic functional model is still not available for this long-known enzyme. Also, the reason for the preference of Ni over metal ions more commonly found in hydrolases such as Zn has remained a mystery. Bioinspired coordination chemistry for new nickel proteins such as ARD is just emerging, and other recently discovered biological active sites such as those in Ni-containing glyoxylase I or Ni-containing peptide deformylase await synthetic model studies. There is no doubt that this area of (bio)inorganic chemistry will see more breakthrough synthetic achievements in the near future, and that biological nickel will remain a great inspiration for synthetic chemists for years to come.

ACKNOWLEDGMENTS FM gratefully acknowledges generous support by the Deutsche Forschungsgemeinschaft (DFG) for our bioinspired work related to the urease active site. JIvdV thanks the Alexander von Humboldt Foundation for a postdoctoral research fellowship.

ABBREVIATIONS acac ACS AHA ARD bdt Bu

acetylacetonate acetyl coenzyme A synthase acetohydroxamic acid acireductone dioxygenase 1,2-benzenedithiolate butyl

Met. Ions Life Sci. 2, 181–240 (2007)


CD CH3SCoM cod CODH Cp depe DFT dmAphaH DMF dmgBF2 dmpe DMPO DMSO dppe ENDOR ESI-MS ESR Et EXAFS H2dsdm HSCoB HYSCORE LMCT MCD MCR Me NBT NHE NiIF430 NiIF430M NiSOD oeibc OTf pdt Ph phen phmi PhTttBu Pr py SCE Stip terpy

231

circular dichroism methylthioethyl sulfonate (methyl coenzyme M) 1,5-cyclooctadiene carbon monoxide dehydrogenase cyclopentadienyl 1,2-bis(diethylphosphino)ethane density functional theory 2-(dimethylamino)phenylhydroxamic acid dimethylformamide (difluoroboryl)dimethylglyoximato 1,2-bis(dimethylphosphino)ethane 5,5-dimethyl-1-pyrrolyl N-oxide dimethylsulfoxide 1,2-bis(diphenylphosphino)ethane electron nuclear double resonance electron spray ionization mass spectrometry electron spin resonance (spectroscopy) ethyl extended X-ray absorption fine structure N,N-dimethyl-N,N-bis(2-sulfanylethyl)ethylenediamine 7-mercaptoheptanoylthreonine phosphate (coenzyme B) hyperfine sublevel correlation spectroscopy ligand-to-metal charge transfer magnetic circular dichroism methyl coenzyme M reductase methyl para-nitro blue tetrazolium chloride normal hydrogen electrode Ni tetrahydrocorphinoid cofactor pentamethylester of NiIF430 nickel-containing superoxide dismutase octaethylisobacteriochlorin triflate trifluoromethanesulfonate 1,3- propanedithiolate phenyl phenanthroline N,N-1,2-phenylenebis(2-sulfanyl-2-methyl)propionamide phenyltris((tert-butylthio)methyl)borate propyl pyridine standard carbon electrode 2,4,6-tris(isopropyl)thiophenolate 2,2,6,2-terpyridine Met. Ions Life Sci. 2, 181–240 (2007)

232


THF tmc tmtaa tsalen

tetrahydrofuran tetramethylcyclam 6,8,15,17-tetramethyldibenzo[b,i][1,4,8,11]tetraazacyclotetradecine N,N-ethylenebis(thiosalicylidene)iminato

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6 Urease: Recent Insights on the Role of Nickel Stefano Ciurli Laboratory of Bioinorganic Chemistry, Department of Agro-Environmental Science and Technology, University of Bologna, Viale Giuseppe Fanin 40, I-40127 Bologna, Italy <[email protected]>

1.

INTRODUCTION: UREASE AND ITS BIOLOGICAL SIGNIFICANCE 2. THE BIOCHEMISTRY OF UREASE 3. STRUCTURAL STUDIES ON BACTERIAL UREASES 3.1. Structures of Urease in the Native State 3.2. Structure of Urease in a Complex with a Substrate Analog 3.3. Structure of Urease in a Complex with a Transition State Analog 3.4. Structures of Urease in Complexes with Competitive Inhibitors 3.5. Structures of Urease Mutants 3.6. Structure of a Metal-Substituted Urease 4. THE STRUCTURE-BASED MECHANISM OF UREASE 4.1. Mechanism Step I: Binding of Urea to the Active Site 4.2. Mechanism Step II: Nucleophilic Attack by the Bridging Hydroxide to Give the Tetrahedral Transition State 4.3. Mechanism Step III: Protonation of the Distal Urea Nitrogen Followed by Release of the First Ammonium Ion



242 243 244 249 255 256 257 263 266 267 268 270 271

242

5.

CIURLI

CONCLUSIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

273 273 273 274

1. INTRODUCTION: UREASE AND ITS BIOLOGICAL SIGNIFICANCE Urea is produced in a large number of vertebrates as the catabolic product for nitrogen-containing compounds. Urease (urea aminohydrolase E.C. 3.5.1.5) is the biological catalyst produced by plants, algae, fungi, and bacteria for the hydrolytic decomposition of urea and release of essential nitrogen [1]. In plants, the primary role of urease is to recycle arginase-derived urea nitrogen during germination [2], although insecticidal activity as a plant defense system has also been reported [3]. The overall hydrolysis of the reaction products causes a pH increase and a consequent release of ammonia, chemical processes that are responsible for the negative side effects of the action of bacterial urease for human health as well as for agriculture. Urease is a virulence factor in human and animal infections of the urinary and gastrointestinal tracts, and is involved in kidney stone formation, catheter encrustation, pyelonephritis, ammonia encephalopathy, hepatic coma, and urinary tract infections [4,5]. Urease is also produced in large amounts (10–15% of total proteins by weight) in Helicobacter pylori, a ureolytic bacterium able to survive in the acidic environment of the stomach by exploiting the pH increase caused by the urease activity, and acting as the major cause of pathologies (including cancer) induced by gastroduodenal infections [6] (see also Chapter 15). In another context, soil urease, present both in living ureolytic bacteria [7] and as extracellular enzyme stabilized by aggregation with clays and humic substances [8,9], degrades urea, a worldwide used soil nitrogen fertilizer [10]. Although necessary for crop nitrogen uptake, the efficiency of soil nitrogen fertilization with urea is severely decreased by the urease activity itself, and large amounts of ammonia nitrogen are lost in the atmosphere, while plant damage occurs by ammonia toxicity and soil pH increase, thereby causing significant environmental and economic problems. Urease was the first enzyme ever to be crystallized, from the plant source Canavalia ensiformis (jack bean) [11], and was also the first protein shown to contain essential nickel ions in the active site [12]. This chapter focuses on the most recent advances in the understanding of the chemistry of nickel in urease, and follows on from the chapter by Andrews, Blakeley, and Zerner in 1988 in Met. Ions Life Sci. 2, 241–278 (2007)

UREASE: RECENT INSIGHTS IN THE ROLE OF Ni

243

the Metal Ions in Biological Systems series [13]. Since then, significant advances have been achieved in the understanding of urease, and several reviews have been published that deal with various aspects of the chemistry and biology of urease [1,4,5,14–17]. This review will take into consideration only the implications on the chemistry of nickel in urease as derived from the crystallographic structural data obtained since the release of the first structure of this protein in 1995 [18] until the most recent days.

2.

THE BIOCHEMISTRY OF UREASE

The urea molecule spontaneously decomposes in water to yield cyanic acid and ammonia, through an elimination process that proceeds with a half-life of ⬃3.6 yr at 38⬚C independently of pH in the 2–12 range [19] (Figure 1A). Considering the large amount of urea continuously released in the environment (for example each human produces ⬃10 kg urea per year [20]), this molecule would rapidly accumulate. The enzymatic decomposition pathway catalyzed by urease causes the formation of ammonia and carbamic acid with kcat /Km approximately 1014 times higher than the rate of the uncatalyzed elimination [21] (Figure 1B), reducing the half-time of urea hydrolysis to few microseconds. The carbamate produced during this reaction spontaneously and rapidly decomposes, at physiological pH, to give a second molecule of ammonia and bicarbonate. The enzyme is localized into the cytoplasm in all sources so far examined [4] with the exception of H. pylori, for which the urease released by autolysis and adsorbed onto the walls of viable cells accounts for ⬃75% of the total activity [22]. Urea is not the only substrate for the enzyme, and hydrolytic activity involving formamide, acetamide, N-methylurea, N-hydroxyurea, N,N⬘-dihydroxyurea, semicarbazide, O (A)

non-catalyzed

C H2N

NH2

ca. 3.6 years

HN

O (B)

C H2N

+ H2O NH2

O

+ NH3

O

urease ca. 1 µs

C

+ NH4+

C H2N

O O

H2O

+ NH3

C HO

O

Figure 1. (A) reaction scheme for the uncatalyzed urea hydrolysis; (B) reaction scheme for the urease-catalyzed urea hydrolysis. Met. Ions Life Sci. 2, 241–278 (2007)

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CIURLI

and different kinds of phosphoric acid amides has been observed [19]. The values of kcat for these alternative substrates is ⬃102–103-fold lower than that observed for the natural substrate urea. Ureases exhibit typical Michaelis–Menten behavior, with no indication of substrate inhibition or allostery [4,5]. The observed values of Km fall in the 1–100 mM range, and are largely independent of pH. On the other hand, k cat and k cat /Km strongly depend on this parameter, and typical bell-shaped pH profiles are observed, which are characterized by pKa values of ⬃6.5 for a general base, which must be deprotonated, and ⬃9.0 for a general acid, which must be protonated. The maximum of activity is detected at pH ⬃7.5–8, with maximal values of k cat and k cat /Km of ⯝3000 s⫺1 and ⯝1000 s⫺1 mM⫺1, respectively [4,5]. Although enzyme inactivation at pH lower than ⬃5 is observed due to irreversible denaturation and loss of the essential nickel ions, a distinct group of ureases possess an optimum pH in the range 2–4.5 [23,24]. The amino acid sequence for these ‘acidic’ enzymes does not differ from that of the ‘neutral’ ureases as far as the active site residues are concerned, and therefore the factors that determine this difference in the pH-dependent activity profiles are still unknown.

3.

STRUCTURAL STUDIES ON BACTERIAL UREASES

Since the first report of the X-ray crystal structure of Klebsiella aerogenes urease (KAU) [18], several refined models of this enzyme [25–30] and the related urease from Bacillus pasteurii (BPU) [31–35] and Helicobacter pylori (HPU) [36] have been released in the Protein Data Bank (PDB), and now the total number sums up to 35 (see Table 1). The available structures are related to the state of the enzyme in its native form with an intact active site, as well as to those of urease mutants, or urease complexed with small molecules. The structures of urease mutants have been determined in order to elucidate the role of the mutated residue in the active site construction or in the catalytic cycle. On the other hand, the structures of urease complexed with small molecules have been determined to elucidate the chemical reactivity of the nickel ions and the surrounding active site residues, and to clarify their role in the catalytic mechanism as well as the mechanism of competitive inhibition of the enzyme. Figure 2 shows the protein architecture for KAU, BPU, and HPU. KAU and BPU are organized in heterotrimers of the type (αβγ)3, with α, β, and γ being three different subunits. The active site is found in the α subunit, giving rise to three active sites per biological unit. On the other hand, in HPU the quaternary structure is built by first organizing the two subunits α and β in a trimer of the type (αβ)3, with an overall structure very similar to the structures of KAU and BPU. In HPU, the α subunit is highly homologous to the α subunit in KAU and BPU, while the Met. Ions Life Sci. 2, 241–278 (2007)


245

Figure 2. (A) Ribbon scheme of the functional oligomer (αβγ)3 of K. aerogenes urease; α subunit: light sea green; β subunit: cyan; γ subunit: blue; (B) ribbon scheme of the functional oligomer (αβγ)3 of B. pasteurii urease; α subunit: yellow; β subunit: orange; γ subunit: red; (C) ribbon scheme of the (αβ)3 oligomer of H. pylori urease; α subunit: pink; β subunit: green; (D) ribbon scheme of the functional oligomer [(αβ)3] 4 of H. pylori urease seen through the ternary axis, front view; (E) ribbon scheme of the functional oligomer [(αβ)3] 4 of H. pylori urease seen through the ternary axis, back view; (F) ribbon scheme of the functional oligomer [(αβ)3] 4 of H. pylori urease seen through the binary axis. For D, E, and F each (αβ)3 unit is shown in different colors: pink, orange, cyan, and yellow. Molecular graphics images were produced using the UCSF Chimera package from the Resource for Biocomputing, Visualization, and Informatics at the University of California, San Francisco (supported by NIH P41 RR-01081) [79]. β subunit corresponds to the fusion of the β and γ subunits found in the KAU and BPU urease. Furthermore, in HPU a higher degree of oligomerization produces the spherically shaped tetramer of trimers [(αβ)3] 4, yielding a biological unit that contains 12 active sites. Such a higher oligomer is not a solid state artifact, but corresponds well to the shape estimated from electron microscopy images of the enzyme immobilized on a support film [37]. The tetramerization of the triangularshaped (αβ)3 unit is believed to be responsible for the acid-resistance of HPU, a peculiar feature that could be important to protect the extracellular enzyme found in the gastric mucosa, through a mechanism defined as ‘intrasupramolecular’. This mechanism implies that the 12 active sites of each functional dodecamer increase the pH locally, thus decreasing the acid-induced loss of the nickel ions from the active site, which would otherwise result in enzyme deactivation. Met. Ions Life Sci. 2, 241–278 (2007)

Enzyme form

Wild-type Apo-form Hisα219Ala Hisα320Ala Cysα319Ala Cysα319Ala Cysα319Ala Cysα319Ala Cysα319Ala Cysα319Asp Cysα319Ser Cysα319Tyr Hisα134Ala Wild-type Lysα217Glu Cysα319Ala Lysα217Cys Lysα217Ala Lysα217Ala Cysα319Ala Lysα217Cys Wild-type Asp α221Ala Hisα219Asn

Enzyme source

K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes

K. aerogenes K. aerogenes K. aerogenes

K. aerogenes K. aerogenes K. aerogenes

PDB Code

1KAU 1KRA 1KRB 1KRC 1FWA 1FWB 1FWC 1FWD 1FWE 1FWF 1FWG 1FWH 1FWI 1FWJ 1A5K 1A5L

1A5M 1A5N 1A5O

Met. Ions Life Sci. 2, 241–278 (2007)

1EF2 1EJR 1EJS

7.5 7.5 7.5

7.5 7.2 7.2

7.5 7.5 7.5 7.5 7.5 6.5 8.5 9.4 7.5 7.5 7.5 7.5 7.5 7.5 7.5 7.5

pHa 2Ni, Water – 2Ni, Water 2Ni, Water 2Ni, Water 2Ni, Water 2Ni, Water 2Ni, Water 2Ni, AHA 2Ni, Water 2Ni, Water 2Ni, Water 1Ni, Water 2Ni, Water – – May 1998 May 1998 May 1998

July 1995 October 1995 October 1995 October 1995 October 1997 October 1997 October 1997 October 1997 October 1997 October 1997 October 1997 October 1997 October 1997 October 1997 May 1998 May 1998 Closed Disordered Closed

Closed Closed Closed Closed Closed Closed Closed Closed Disordered Disordered Closed Open Disordered Closed Closed Disordered

2.50 2.00 2.00

2.00 2.40 2.50

2.20 2.30 2.50 2.50 2.00 2.00 2.00 2.00 2.00 2.00 2.00 2.00 2.00 2.20 2.20 2.20

99 100 96

90 100 100

94.6 98 98 98 94 97 96 96 96 97 92 91 93 96 92 93

17.2 16.1 18.5

19.4 16.7 18.1

18.2 19.0 17.9 18.0 16.9 16.9 16.8 16.9 20.4 17.1 17.6 17.9 17.0 17.3 17.8 18.6

Data Flap status Resolution completeness R-value (Å) (%) (%)

298 2Mn, Water March 2000 Closed 298 2Ni, Water September 2000 Disordered 298 2Ni, Water September 2000 Closed

298 – 298 Formate 298 Formate

298 298 298 298 298 298 298 298 298 298 298 298 298 298 298 298

T Non-protein Release date (K) b ligands

Table 1. Selected structural information for urease structures currently available in the Protein Data Bank

[29] [30] [30]

[28] [28] [28]

[18] [25] [25] [25] [26] [26] [26] [26] [26] [26] [26] [26] [27] [26] [28] [28]

Reference

246

b

a

K. aerogenes K. aerogenes K. aerogenes K. aerogenes K. aerogenes B. pasteurii B. pasteurii B. pasteurii B. pasteurii B. pasteurii B. pasteurii H. pylori H. pylori

Hisα219Gln Hisα320Asn Hisα320Gln Wild-type Wild-type Native Native Native Native Native Native Native Native

pH of crystallization. Temperature for data collection.

1EJT 1EJU 1EJV 1EJX 1EJW 2UBP 1UBP 3UBP 4UBP 1IE7 1S3T 1E9Y 1E9Y

7.5 7.5 7.5 7.5 7.5 6.8 6.8 6.8 6.8 6.8 6.8 6.5 6.5

298 298 298 100 298 100 100 100 100 100 100 100 100

2Ni, Water 2Ni, Water 2Ni, Water 2Ni, Water 2Ni, Water 2Ni, Water 2Ni, BME 2Ni, DAP 2Ni, AHA 2Ni, PHO 2Ni, H3BO3 2Ni, Water 2Ni, AHA

September 2000 September 2000 September 2000 July 2003 November 2003 November 1999 March 1999 December 1999 March 2000 April 2001 April 2004 November 2001 November 2001

Closed Disordered Disordered Disordered Closed Open Open Closed Open Open Open Closed Open

2.00 2.00 2.40 1.60 1.90 2.00 1.65 2.00 1.55 1.85 2.10 3.00 3.00

96 91 99 93 95 96.7 98.7 99.9 99.5 99.3 100 95.3 92.5

17.3 17.3 18.1 17.2 14.4 16.0 16.0 15.8 19.0 19.0 17.8 21.2 21.3

[30] [30] [30] unpublished unpublished [31] [32] [31] [33] [34] [35] [36] [36]

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In addition to the structures of KAU, BPU and HPU, a paper reporting the preliminary crystal structure determination of jack bean urease (JBU) has also appeared in the literature [38]. The resolution is rather low (4 Å), but sufficient to identify the overall structural architecture of the molecule as built from the dimerization of two trimers, with each trimer formed using three α subunits. Each α subunit is highly homologous to the αβγ trimer found in KAU and BPU, so the α3 trimer is structurally very similar to the (αβγ)3 trimer of KAU and BPU. This structure of JBU has not been deposited in the PDB and therefore no refined model can be discussed here. The secondary and tertiary structure of KAU, BPU and HPU are very similar. The α subunits of the three proteins make up the body of the protein trimer and consist of an (αβ) 8 barrel domain and a β -type domain. The β subunits of KAU and BPU are located on the external surface of the trimer and are mainly composed of β strands, with an additional α helix present in BPU. The γ subunits of KAU and BPU consist of αβ domains located on top of each pair of α subunits, thereby favoring their association in a trimer. A peculiar feature of the γ subunit in BPU is the presence of a carbamylated N-terminus, well packed between the γ and the α subunit, and actually contributing to the stability of this aggregate through strong H bonds that involve the terminal carbamate group and few sidechains of the α subunit. This carbamate group is deeply buried and shielded from the protein surface. Therefore, this post-translational chemical modification must occur during the assembly of the heterotrimer and not as the result of a subsequent action by the chaperones involved in the carbamylation of the active site lysine (see Chapter 14). The origin and presence of this structural feature is possibly important in order to understand the process of assembly of the functional enzyme from the single subunits. In HPU, the β subunit consists of two domains: one domain corresponds to the β subunit of KAU and BPU and features very similar secondary and tertiary structure, with the same α helix as found in BPU but also with an additional fragment of 11 residues, which forms a terminal loop believed to be critically involved in the tetramerization of the (αβ)3 trimer to yield the ball-shaped HPU functional unit. The second domain of the β subunit of HPU is essentially identical to the γ subunit of KAU and BPU. A long protein fragment, stretching ⬃30 Å, connects the two domains of the β subunit of HPU. A peculiar feature of the structures of native KAU, BPU and HPU is the conformation of the helix–turn–helix motif (residues 311–340 in the BPU structure) of the α subunit, flanking the active site cleft where the two nickel ions are found. This motif is characterized by high mobility, a feature thought to be important for the regulation of the flux of reactants and products towards and from the active site cavity, thereby acting as a flap able to open or close. Indeed, this motif is found in a more ‘open’ conformation in the structure of native BPU, while it can be defined as existing in a ‘closed’ conformation in native KAU and native HPU (Figure 3). As discussed below, the conformation of this flap can change, and it is Met. Ions Life Sci. 2, 241–278 (2007)


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Figure 3. Chain trace of the active site flap of native K. aerogenes urease (KAU, cyan), B. pasteurii urease (BPU, yellow), and H. pylori urease (HPU, red), highlighting the ‘open’ and ‘closed’ conformations. The nickel ions in the active site are shown as spheres of arbitrary radius. Molecular graphics images were produced using the UCSF Chimera package from the Resource for Biocomputing, Visualization, and Informatics at the University of California, San Francisco (supported by NIH P41 RR-01081) [79].

related to the presence, in the active site, of the reactants, intermediates, or product of the catalytic hydrolysis of urea, and of molecules of inhibitors.

3.1.

Structures of Urease in the Native State

Four different structures of native KAU are available (PDB codes 1KAU, 1FWJ, 1EJW, and 1EJX, see Table 1 and Figure 4). They reveal a substantial identity of the arrangement of the active site residues, but also small differences in the presence and location of the water molecules. These are important, as water is one of the reactants of the catalyzed reaction of urea hydrolysis. In native KAU, two Ni atoms are held at a distance of 3.5–3.7 Å by the bridging carboxylate group of the carbamylated Lysα217 residue, bound to Ni(1) through Oθ1 and to Ni(2) through Oθ2. Ni(1) is further bound to Hisα246 Nδ and to Hisα272 Nε, while Ni(2) is bound to Hisα134 Nε, to Hisα136 Nε, and to Aspα360 Oδ1. In the first model proposed (PDB code 1KAU), Ni(1) appeared to be tri-coordinated, while Ni(2) was penta-coordinated in a distorted trigonal bipyramidal geometry (or distorted square pyramidal, with Hisα136 Nε as the apical ligand) [18]. The fifth ligand Met. Ions Life Sci. 2, 241–278 (2007)

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Figure 4. Ball-and-stick drawing of the crystallographic structural models for the active site obtained for K. aerogenes urease in the native state. The PDB codes are indicated. H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine, R ⫽ arginine, W ⫽ solvent molecules. The nickel ions are represented as black spheres.

to Ni(2) was assigned as a well-ordered water molecule (W2) proposed to act as the catalytic nucleophile, while Hisα320 Nε was described as deprotonated (neutral form) and acting as the general base in the catalysis [18]. Later, the same authors proposed that Hisα320 Nε is protonated at pH 7.5 and could act as the general acid [14,27], a proposal that somehow contrasted with its pKa ⫽ 6.5 [39,40]. In native KAU, Hisα219, known to be involved in substrate binding [40], is placed at 3.1 Å from Ni(1), pointing toward the active site cavity with its protonated Nε atom: this feature suggested that it could act as hydrogen bonding donor to the oxygen of urea, thereby helping to position and polarize the substrate [25]. However, in this structure no hydrogen bonding partner was found, as Hisα219 pointed to an Met. Ions Life Sci. 2, 241–278 (2007)


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empty coordination site of Ni(1). The lack of light, non-protein ligands, possibly water molecules or hydroxide ions, in the active site of KAU, was probably due to the relatively poor data quality, and indeed the electron density suggested the presence of other water positions that were not well refined [18]. The proposed structural model was in disagreement with previous spectroscopic data on KAU [41], JBU [41–47], and BPU [48], which indicated the presence of hexa- or pentacoordinated slightly distorted octahedral Ni ions. Later, the same authors released another structural model (PDB code 1FWJ, see Table 1 and Figure 4) in which the nickel coordination sites where filled with three solvent-derived oxygen-based ligands [26]. In particular, a nickel bridging ligand (W B) and two water molecules were bound to Ni(1) and Ni(2) (W1 and W2, respectively) at 2.0–2.2 Å, producing a distorted square-pyramidal penta-coordinate Ni(1) and a pseudo-octahedral hexa-coordinate Ni(2), coordination numbers and geometries fully in agreement with the spectroscopic data. In this refined model, the distances between the three solvent molecules (2.0–2.5 Å) were considered too short to allow them to be present simultaneously, and disorder of the water molecules was proposed to be present in the active site [26]. More recently, two additional structures have been determined for native KAU, with data collected at 100 K (PDB code 1EJX) and at 298 K (PDB code 1EJW), which confirmed the presence of W B, W1 and W2 with no substantial difference between the structures determined at the two temperatures. A remarkable feature of 1EJX is the anomalous distance of 2.5 Å between Ni(2) and W2, while no significant alteration of the distances between the three solvent molecules is observed with respect to the 1FWJ model. In all three models 1FWJ, 1EJW and 1EJX, a fourth water molecule is found in the active site cavity (W3). This water molecule is at the center of a complex H-bonding network involving W1, W2 and W B as well as the carbonyl oxygen of the nearby Ala α363 and the Nε atom of His α320. This latter residue is located on the active site flap of KAU, found in the ‘closed’ conformation in all models of the native form of the enzyme except for 1EJX, in which this motif lacks the central fragment of 11 residues, probably because of disorder. As such, His α320 is placed relatively close to the active site core, and could in principle assume the role of a proton shuttle in the catalytic mechanism. The structure of apo-urease from K. aerogenes (PDB code 1KRA, Table 1), in which the Ni ions were chemically removed by lowering the pH in the presence of chelating agents, features a de-carbamylated Lysα217 residue [25]. All other α-carbon atoms show a very small root-mean-square deviation (0.2 Å) with respect to the holo-enzyme, indicating an overall rigidity of the protein, and in particular of the active site. This suggests that the architecture of urease is designed to bind the Ni ions in a well-defined mode without having to undergo conformational rearrangements that might destabilize the functioning enzyme and increase the Ni dissociation equilibrium constant. Met. Ions Life Sci. 2, 241–278 (2007)

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The structure of another urease in the native state, from B. pasteurii (PDB code 2UBP, Table 1 and Figure 5) has also been reported [31]. In BPU, the electron density in the active site is very well defined, providing a clear picture of the Ni coordination. The two metal ions are bridged by the carboxylate group of the carbamylated Lys α220, while Ni(1) is further coordinated by His α249 Nδ and His α275 Nε , and Ni(2) is bound to His α137 Nε , His α139 Nε , and Asp α363 Oδ1. The Ni ions are separated by 3.7 Å. By analogy with the refined structures of native KAU, four water/hydroxide molecules, three of which directly bound to the metal ions, where modeled in the active site of BPU. One of these (W B) symmetrically bridges the two Ni ions, whereas the other two (W1 and W2) complete the coordination polyhedron around the Ni ions. A fourth water molecule, W3, strongly interacts with W B (at 2.3 Å), W1 (at 2.2 Å), and W2 (at 2.4 Å), through hydrogen bonds. The angles formed by Ni(1,2)–W(1,2,B)– W(3) are very close to tetrahedral. W B is at hydrogen bonding distance from Asp α363 Oδ2 (at 2.5 Å). W3 is at the center of additional possible multiple hydrogen bonding interactions, being 3.3 Å from Asp α363 Oδ2 and at 3.0 Å from the oxygen atom of a sulfate ion found in the cavity. The sulfate ion is located between the four water/hydroxide cluster and the nearby Argα339, in a position occupied, in native KAU, by the imidazole ring of His α323 (His α320 in native KAU). Argα339 extends from the very bottom of the active site cleft, and forms a strong salt bridge with the sulfate ion, which in turn forms a hydrogen bond with His α323 Nε (at 2.7 Å). The presence of sulfate is probably due to its high concentration in the crystallization buffer. The assignment of the protonation state of W1 and W2 as water molecules was suggested [31] by the value of the first dissociation constant for Ni(H2O) 2⫹ 6 (pKa ⫽ 10.6) [49]. In water-bridged bimetallic complexes, the first pKa for the bridging water decreases significantly to very acidic values, while the pKa for hydroxide deprotonation is slightly lower than the pKa of the first ionization of a water bound to a single Ni ion [49]. Therefore, the estimated pKa for the deprotonation of the Ni bridging hydroxide (⬃9–10) suggests that, at pH ⫽ 8.0, WB is in the hydroxo form. The distances between WB, W1, and W2 in native BPU are as short as those found in native KAU (2.1–2.2 Å). However, for native BPU this was not explained as due to partial occupancy as in the case of KAU, but rather as a consequence of the presence of a four-centered hydrogen bonding network involving a proton located in the center of the tetrahedron constituted by W1, W2, W3 and WB [31]. Moreover, W1 is at 2.9 Å from Hisα222 Nε, which is protonated and acts as a hydrogen bonding donor, as deduced from the interaction of Hisα222 Nδ with the peptide NH group of Aspα224 (at 2.9 Å). In contrast, W2 forms a strong hydrogen bond with Alaα170 O (at 2.9 Å), which acts as hydrogen bonding acceptor. The positions of conserved amino acid residues not involved in Ni binding, but thought to be important in the catalytic mechanism (Ala α170, Hisα222, Glyα280, Cysα322, Hisα323, Hisα324, Argα339, and Ala α366) (BPU consensus sequence) are Met. Ions Life Sci. 2, 241–278 (2007)


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Figure 5. Ball-and-stick drawing of the crystallographic structural models for the active site obtained for B. pasteurii urease (PDB code 2UBP) and H. pylori urease (PDB code 1E9Z) in the native state. H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine, R ⫽ arginine, W ⫽ solvent molecules. The nickel ions are represented as black spheres.

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largely invariant in BPU and KAU, except for Cysα322, Hisα323, and Hisα324. In particular, in native BPU, Hisα323 is displaced by ⬃5 Å away from W3 with respect to the position of the homologous residue His α320 in native KAU, where Hisα320 is located at hydrogen bonding distance to W3, while in BPU the corresponding interaction between Hisα323 and W3 is hindered by the presence of the sulfate ion. This difference is due to the fact that, in native BPU, the helix–turn–helix motif lining the active site cleft in the α subunit is found in the ‘open’ conformation in BPU as opposed to being ‘closed’ in KAU. The opening of the flap in native BPU also induces a large shift of the position of Cysα322 (ca. 4 Å) and of Hisα324 (ca. 7 Å) with respect to the position of the homologous residues Cys α319 and Hisα321 in native KAU. Furthermore, in KAU the carbonyl oxygen of Ala α363 is at hydrogen bonding distance from W2, whereas, when the flap is open as in BPU, the corresponding residue Ala α366 is rotated away, at a distance (5.1 Å from W2) that does not allow formation of hydrogen bonds. The position of the side chain of Argα339, stretching from the very bottom of the cleft toward the active site Ni ions, is not influenced as much as for Cysα322, Hisα323, and Hisα324 by the flap opening. The latter induces a shift of the terminal guanidinium group of only ca. 1 Å away from the Ni ions with respect to the position of the homologous Argα336 in KAU. The case of native HPU (Figure 5) deserves special attention. Indeed, large differences occur in this structure compared with KAU and BPU, and these will be discussed here, with the caveat that such structure was solved with much lower resolution (3.00 Å instead of 1.55 Å for BPU and 2.00 Å for KAU). The most striking difference is represented by the bridging mode of the carbamylated Lys β219, found here to bridge the two nickel ions using only one of the two terminal oxygen atoms, with a Ni–O distance of 2.7 Å. The second oxygen atom interacts even more weakly with Ni(1) at a distance of 2.8 Å, while it is too far from Ni(2) to be involved in any bonding interaction (3.4 Å). This is caused by an apparent rotation of the terminal –NH–CO⫺2 moiety along the N–C bond by about 90⬚ with respect to the position found in BPU and KAU. The distances between the nickel ions and the coordinating residues are very large (ranging between 2.6 and 2.7 Å) as compared to those found in KAU and BPU (2.0–2.1 Å) with the exception of the interaction between Ni(2) and His β138 (2.3 Å) and Asp β362 (2.2 Å). In addition, the orientation of the histidine imidazole rings around the metal ions is quite unusual, as is evident from Figure 5. Even more strikingly, the Ni–Ni distance is reduced to the very short value of 2.1 Å. No solvent-derived oxygen atom is found bridging the metal ions, and a single water molecule is found in the active site, weakly interacting with Ni(1) at 2.6 Å. In this situation, the Ni ions would be tetra-coordinated. These observations are too anomalous with respect to the situation found in several structures of the other two bacterial ureases, and certainly deserve more attention and possibly a higher resolution model.

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3.2. Structure of Urease in a Complex with a Substrate Analog The determination of the structure of urease with nonreactive analogs of urea could in principle help in the elucidation of the binding mode of the substrate, the first step of the catalytic mechanism. Boric acid, B(OH)3, known to be a competitive inhibitor of urease from jack bean [50], Proteus mirabilis [51], and Klebsiella aerogenes [52], is isoelectronic with urea, has the same triangular shape and dimension, and, like urea, has a neutral charge, and can thus be considered a good substrate analog. Therefore, the structure of BPU crystallized in the presence of B(OH)3 (PDB code 1S3T, Table 1 and Figure 6) has been determined [35]. The refined model confirms the rigidity of the protein scaffold, with an overall backbone RMSD of only 0.2 Å from the structure of the native enzyme. The helix–loop–helix flap flanking the active site channel is in the open conformation, as found in the native enzyme [31]. In the structure, a molecule of B(OH)3 is symmetrically placed between the Ni ions, substituting W1, W2, and W3 and leaving in place the bridging hydroxide while not perturbing the Ni–Ni distance (3.7 and 3.6 Å in the native and B(OH)3-inhibited forms, respectively).

Figure 6. Ball-and-stick drawing of the crystallographic structural model for the active site obtained for B. pasteurii urease complexed with boric acid B(OH)3. H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine, R ⫽ arginine, W ⫽ solvent molecules. The nickel ions are represented as black spheres.

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Two inhibitor oxygen atoms are bound to the Ni ions (at 2.2 and 2.1 Å) while the third oxygen points towards the cavity opening, away from the Ni ions. The geometry and coordination number of the Ni ions do not change significantly upon boric acid binding. The protonation state of the inhibitor is well established as the neutral B(OH)3 form from a detailed analysis of the surrounding H-bonding network, the role of neighboring residues capable to stabilize an unprecedented binding mode of B(OH)3 to a dinuclear metal center. This model constitutes a significant basis for any mechanistic proposal for urease catalysis, and in particular for the first step of the mechanism, the binding of the substrate.

3.3. Structure of Urease in a Complex with a Transition State Analog In order to propose a reaction mechanism for enzymatic urea hydrolysis, the knowledge of the structure of the transition states or the intermediates of the reaction is essential. The very short lifetime of such chemical moieties prevents this information to be attained using X-ray crystallography. However, this technique could exploit the lack of reactivity of analogs of the transition state. A good transition state analog for the enzymatic hydrolysis of urea is the tetrahedral diamidophosphate (NH2)2PO⫺2 (DAP). Evidence that, in the presence of phenylphosphorodiamidate (NH2)2PO(OPh) (PPD), the actual inhibitor of urease is DAP, produced by the enzymatic hydrolysis of PPD to yield phenol, was suggested by kinetic experiments [53]. BPU was therefore crystallized in the presence of PPD, and the structure of the resulting complex was obtained (PDB code 3UBP, Table 1 and Figure 7) [31]. This structure reveals that in the active site the positions of all Ni-bound protein residues do not significantly differ from the structure of native BPU. A tetrahedral molecule, which replaced the cluster of four water/hydroxide molecules found in native BPU, was found at the active site. Clearly, the bound inhibitor was not the PPD molecule, due to the absence of electron density accounting for an aromatic ring, and therefore the electron density was modeled with DAP. A specific hydrogen bonding pattern stabilizes the inhibitor and directs its orientation in the cavity, allowing the assignment of the Ni-bound DAP atoms to either oxygen or nitrogen. DAP is bound to Ni(1) and to Ni(2) through one oxygen and one nitrogen atom, respectively. The second DAP oxygen symmetrically bridges the two nickel ions, while the second DAP nitrogen atom points away towards the cavity opening. The Ni(1)-bound oxygen atom receives a hydrogen bond from Hisα222 Nε, while the Ni(2)-bound DAP NH2 group forms two hydrogen bonds, donated to the carbonyl oxygen atoms of Alaα170 and Alaα366. The Ni-bridging DAP oxygen atom is at a hydrogen bonding distance from Aspα363 Oδ2 atom. In DAPinhibited BPU, the flap is in the closed conformation, essentially identical to that found in native KAU. This conformation allows the formation of two additional Met. Ions Life Sci. 2, 241–278 (2007)


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Figure 7. Ball-and-stick drawing of the crystallographic structural model for the active site obtained for B. pasteurii urease complexed with diaminophosphate (DAP). H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine, R ⫽ arginine, W ⫽ solvent molecules. The nickel ions are represented as black spheres.

hydrogen bonds between the distal DAP NH2 group and Alaα366 O (found in a different conformation with respect to that of native BPU) and Hisα323 Nε. The latter residue is found in a position analogous to that observed in native KAU.

3.4.

Structures of Urease in Complexes with Competitive Inhibitors

Several classes of molecules (diphenols, quinones, hydroxamic acids, phosphoramides, and thiols) have been tested as urease inhibitors in medicine and agriculture [4,5]. However, the efficiency of the presently available inhibitors is low, and negative side effects on humans and on the environment [4,5,54] have been reported. A structure-based molecular design approach for the discovery of new and efficient ureases relies upon the elucidation of the structure and reactivity of the active site residues. The highly conserved amino acid sequences of all known ureases [5] and the constant presence of two Ni ions and of their ligands in the active sites infer a common chemical behavior. To this end, several structures of BPU complexed with competitive inhibitors other than those described above, which acted as substrate or transition state analogs, have been determined. Met. Ions Life Sci. 2, 241–278 (2007)

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Figure 8. Ball-and-stick drawing of the crystallographic structural model for the active site obtained for B. pasteurii urease complexed with β -mercaptoethanol (BME). H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine, R ⫽ arginine, C ⫽ cysteine. The nickel ions are represented as black spheres.

The structure of BPU inhibited with β -mercaptoethanol (BME) [31,32] (PDB code 1UBP, Table 1, Figure 8) clearly shows the presence of the inhibitor bound to the Ni ions in the active site. The binding of the inhibitor does not modify the arrangement of the surrounding protein residues, a further evidence of the substantial rigidity of the environment of the metal ions. In the active site of the inhibited enzyme, the presence of the BME molecule has caused the expulsion of all four water-based molecules from the active site. The sulfur atom of BME has substituted the WB hydroxide, suggesting that this bridge is labile and that the dinuclear metal center can sustain such substitution. The direct binding of the thiol functionality to the nickel ions is consistent with charge-transfer transitions observed in the near-UV [42] and magnetic circular dichroism (MCD) [47] spectra of urease in the presence of thiolate competitive inhibitors. Magnetic susceptibility and variable-temperature MCD spectra of BME-inhibited urease also indicate that the Ni centers are strongly anti-ferromagnetically exchange-coupled, in agreement with the structural information [46,47]. This observation was further supported by EXAFS [41,45]. The nickel-bridging S atom of BME is at a hydrogen bonding distance from the carbonyl oxygen of Alaα170, and this could imply that the thiol functionality, and not the thiolate, is acting as a ligand. An unexpected feature of the BME binding Met. Ions Life Sci. 2, 241–278 (2007)


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mode is the chelating interaction involving the terminal OH group of the inhibitor, coordinated to Ni(1). This contact is assisted and stabilized by a strong hydrogen bond (at 2.4 Å) donated to the carbonyl oxygen of the conserved Glyα280. Figure 8 shows that, in BME-inhibited BPU, a second molecule of inhibitor is involved in a mixed disulfide bond with Cysα322. This residue is located on the mobile flap, found to be open in BME-inhibited BPU, and it has been proposed to play a significant role in the catalytic process [55,56], even though it is not fully conserved, being replaced in some cases by a threonine residue [57,58]. In addition to acting as a gate to the active site cavity, this flap could take part in the assembly of the dinickel center from apo-urease, for example through the conserved Cysα322, acting, in this case, as a Ni shuttle. The BME molecule forming the mixed disulfide with Cysα322 is further involved in a hydrogen bond between its OH group and the carbonyl oxygen atom of Alaα366, positioned on a neighboring loop (Figure 8). This interaction reduces the flexibility of the flap, and the resulting network seals the entrance to the active site by steric hindrance. In summary, the X-ray structure of BME-inhibited BPU reveals that inhibition occurs by targeting enzyme sites that both directly (the metal centers) and indirectly (the cysteine side chain) participate in substrate positioning and activation. This double inhibition mode is likely to be involved in all examples in which a thiol acts as urease inhibitor [52,55]. Another class of inhibitors of urease is constituted by hydroxamic acid derivatives [52,59,60]. The structures of the complex between acetohydroxamic acid (AHA) and KAU mutant urease (Cysα319Ala) [26], BPU [33], and HPU [36] have been described (PDB codes 1FWE, 4UBP, and 1E9Y respectively, Table 1 and Figure 9). The main structural features of the active site residues are essentially identical to the native forms, and they are supporting, once more, the rigidity of the active site. All three models concur in consistently indicating the binding mode of the inhibitor to the nickel ions in the active site: the negatively charged hydroxamate OB oxygen bridges the two Ni ions, while the carbonyl oxygen of AHA is further bound to Ni(1) at 2.0 Å, in a chelate mode. The binding of the inhibitor to the metal center is assisted by two strong H bonds: the carbonyl AHA O atom receives an H bond from the Hisα222 NεH proton, and the AHA-NH group interacts with the Oδ2 atom of the Ni(2)-bound Aspα363 residue in KAU and BPU, while the latter forms an H-bond with the bridging hydroxamate OB oxygen in HPU. In the case of BPU, the Aspα363 residue undergoes a rotation about the Cβ –Cγ bond by 35⬚ with respect to its conformation in native BPU. No water molecules are found in the vicinity of the metal ions. A pronounced asymmetry was reported for the Ni-bridging atom (Ni(1)–OB ⫽ 2.6 Å; Ni(2)–OB ⫽ 1.8 Å), while in the case of BPU the bridge is symmetric (Ni–OB distances of 2.0 Å). The case of AHA-inhibited HPU is again peculiar because the structure confirms the heretical structural features compared with the ‘canonical’ KAU and BPU, but supports the same type of inhibitor binding mode to the metal atoms in Met. Ions Life Sci. 2, 241–278 (2007)

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Figure 9. Ball-and-stick drawing of the crystallographic structural model for the active site obtained for urease from K. aerogenes (1FWE), B. pasteurii (4UBP) and H. pylori (1E9Y) complexed with acetohydroxamic acid (AHA). H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine. The nickel ions are represented as black spheres. Met. Ions Life Sci. 2, 241–278 (2007)


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the active site. The distances around the Ni ions are still too large to be chemically reasonable, but are certainly closer to the expected values (Ni-Ni: 3.0 Å, Ni-NHis: 2.2–2.3 Å, Ni-OLys: 2.4–2.5 Å). The bridging mode of the carbamylated lysine confirms the apparent rotation along the N–C bond as observed in the native structure. In the structure of AHA-complexed HPU, the conformation of the flap is found in the ‘open’ state as opposed to the ‘closed’ state observed for the structure of native HPU obtained in the absence of AHA in the crystallization buffer. Therefore, a clear conformational change in the HPU active site flap occurs upon inhibitor binding. This is in contrast with what is observed in the case of BPU, for which the flap is open both in the native and in the AHA-inhibited state, while in the case of KAU the flap is highly disordered, and its electron density was not modeled. The apparent higher flexibility of the active site flap observed in HPU was claimed to be due to few residue substitutions in the region acting as the fulcrum of this conformational change [36]. An additional competitive inhibitor for urease is phosphate [61]. Phosphate competitively inhibits KAU in the pH range 5.0–7.0. The inhibition is weak and not purely competitive at pH ⬎ 7.0, while the enzyme is inactive at pH ⬍ 5.0, probably because of the loss of the nickel ions. Within the pH range 5.0–7.0 the inhibition is pH-dependent, with values of Ki increasing from ⬃0.1 mM at pH 5 to ⬃50 mM at pH 7, indicating a progressive decrease of the inhibitory capability. Within this pH range, pKi exhibits a slope of (–1) from pH 5.0 to 6.3, and a slope of (–2) from pH 6.3 to 7.0. These results suggest that phosphate inhibition of urease requires the protonation of an ionizable group with pKa1 ⫽ 6.3 [52]. Similar results were more recently obtained for JBU [62], where the pH-dependence of phosphate inhibition was investigated in the range 5.8–8.1. It was shown that phosphate is a competitive inhibitor in the pH range 5.8–7.5, with Ki values increasing from 0.53 mM at pH 5.8 to 123 mM at pH 7.5. Similar to KAU, the slope of pKi versus pH was (–1) in the pH range 5.8–6.5 and (–2) in the range 6.5–7.5. These results revealed the existence of two ionizable groups having dissociation constants corresponding to pKa1 ⫽ 6.5 (similar to the value of 6.3 observed for KAU) and pKa3 ⫽ 7.2. At pH higher than 7.6 there was no competitive inhibition. Overall, the comprehensive analysis of these data leads to the conclusion that phosphate inhibition of urease involves at least two protonation sites, with pKa values of ⬃7.2 and 6.5. Several different inhibition mechanisms can therefore be proposed, depending on whether these values of pKa are associated to the inhibitor or to active site residues. The structure of BPU crystallized in the presence of phosphate (PHO) could in principle provide a rationale for the kinetic data on phosphate inhibition [34]. The overall structure (PDB code 1IE7, Table 1 and Figure 10) is very similar to that of native BPU with an overall RMSD of only 0.14 Å. Similarly to the other structures of BPU, the active site flap is characterized by large temperature factors, indicating conformational flexibility, and it adopts the ‘open’ conformation as in the native state. Met. Ions Life Sci. 2, 241–278 (2007)

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Figure 10. Ball-and-stick drawing of the crystallographic structural model for the active site obtained for B. pasteurii urease complexed with phosphate (PHO). H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine, R ⫽ arginine. The nickel ions are represented as black spheres.

The arrangement of the protein ligands around the Ni ions in the PHO–BPU adduct is essentially identical to that observed in native BPU. The difference is constituted by the presence of a tetrahedral molecule of phosphate, bound to the binuclear nickel center through three atoms. The Ni-bridging phosphate oxygen atom is symmetrically placed between the two metal ions (at 1.9 and 2.0 Å to Ni(1) and Ni(2), respectively). The OB atom of phosphate replaces the bridging hydroxide molecule (WB) present in native BPU [31]. Two of the other phosphate oxygen atoms coordinate Ni(1) and Ni(2) at ligand distances of 2.4 Å, while the fourth (‘distal’) phosphate oxygen points towards the cavity opening, away from the two nickel ions. As a result of phosphate binding, Ni(1) is penta-coordinate, while Ni(2) is hexa-coordinate. The H-bonding network, which involves the inhibitor atoms and the active site residues, provides a clear definition of the protonation state of the bound phosphate at the crystallization pH of 6.3. The Ni(1)-bound phosphate oxygen atom must be deprotonated, as it receives a strong H-bond from the protonated Hisα222 Nε (at 2.6 Å). As in all urease structures, the protonation state of the latter is dictated by the interaction of Hisα222 Nδ with the peptide NH group of Aspα224. The Ni(2)bound phosphate oxygen atom must be protonated, as it donates a strong H-bond to the carbonyl oxygen atom of Alaα170 (at 2.8 Å). The distal phosphate oxygen Met. Ions Life Sci. 2, 241–278 (2007)


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must be protonated, as it is within H-bonding distance (3.0 Å) of the carbonyl oxygen of Alaα366. This distal phosphate oxygen also receives an H-bond from a water molecule (at 2.8 Å), which additionally interacts with Argα339 through a strong charge-assisted H-bond. The bridging phosphate oxygen must also be protonated at this pH, as it forms an H-bond (at 2.8 Å) with Aspα363 Oδ2. Thus, the Ni-bound phosphate moiety bears three hydrogen atoms and is formally bound as the neutral phosphoric acid H3PO4. These observations suggested an interpretation of the pH-dependence of phosphate inhibition [34]. In the pH range 5–6.5, the most active form of the phosphate inhibitor is H3PO4. The pKa observed at ⬃6.5 was assigned to the loss of the proton shared between phosphate and Aspα363 Oδ2, and corresponds to the production of the H2PO⫺4 species, existing in the pH range 6.5–7.2. Loss of the proton shared between phosphate and Alaα170 O was suggested to correspond to the ionization occurring at pH 7.2, which then involves the formation of the HPO42⫺ species, a form of phosphate unable to inhibit urease [34].

3.5. Structures of Urease Mutants Several mutants of KAU have been structurally investigated in order to rationalize their enzymatic behavior and to establish the role of the mutated residues in the building and functioning of the active site. The importance of the carbamylated lysine for the urease active site building and integrity was investigated through the structural determination of the KAU mutants Lysα217Glu (PDB code 1A5K, Table 1), Lysα217Ala (PDB code 1A5M, Table 1), and Lysα217Cys/Cysα319Ala (PDB code 1A5L, Table 1) [28]. The structures of these mutants reveal the complete absence of bound Ni ions, with the Glu, Ala, and Cys side chains substituting the carbamylated lysine not being able to support the formation of the bimetallic active site. However, all three mutants could be ‘chemically rescued’ by adding small organic acids, such as formic acid, and Ni ions, a procedure that yielded the active site containing both nickel ions. The structures of the Lysα217Ala–formate–Ni complex (PDB code 1A5N, Table 1, Figure 11) and Lysα217Cys/Cysα319Ala–formate–Ni complex (PDB code 1A5O, Table 1) reveal the presence of a dinuclear Ni center bridged by formate instead of the carbamate group of Lysα217 as in wild type KAU [28]. Only one water molecule (W502) was refined in a position close to Ni(2). The mutation of Hisα134, one of the two histidine ligands to Ni(2), to the corresponding alanine residue yields the catalytically inactive mutant Hisα134Ala [40]. The structure of the mutant reveals an enzyme lacking Ni(2) and featuring only Ni(1) in the active site (PDB code 1FWI, Table 1, Figure 11) [27]. The Lys α217* is still carbamylated, and the position of all active site residues (except for the missing Hisα134) highlights again the substantial rigidity of the structure. The Ni(1) ion is octahedrally bound to the same protein ligands as in wild-type urease, in addition to three water molecules. Met. Ions Life Sci. 2, 241–278 (2007)

Figure 11. Ball-and-stick drawing of the crystallographic structural model for the active site obtained for K. aerogenes urease mutants (see Table 1). H ⫽ histidine, K ⫽ lysine, D ⫽ aspartate, A ⫽ alanine, R ⫽ arginine. The nickel ions are represented as black spheres.

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Two residues not directly bound to the nickel ions in KAU, but well integrated in the active site and interacting with the Ni-bound water molecules through H-bonds are Hisα219 and Hisα320. Hisα219 has a direct or indirect role for Hisα219 in substrate binding, as indicated by the much lower affinity for the substrate urea (Km increases ⬃103-fold) for the Hisα219Ala mutant [40]. On the other hand, Hisα219 is not involved in acid/base catalysis, as indicated by the similar rate of enzymatic urea hydrolysis of the same mutant as well as Hisα219Asn and Hisα219Gln [30, 40]. The structure of this mutant (PDB code 1KRB, Table 1, Figure 11) shows substantial identity with native KAU, including the position of the two water molecules in the vicinity of the Ni ions, except with the obvious absence of Hisα219 and the presence, in its place, of Ala α219 [25]. The importance of Hisα219 in the placement of the water molecule bound to Ni(1), is revealed by the absence of this ligand in the structure of the mutant. On the other hand, for the Hisα219Asn and the Hisα219Gln mutants the decrease in affinity is ⬃10-fold less marked. This could be due, in principle, to the fact that the latter two mutants retain H-bonding capability, and indeed this is observed in their crystal structures (PDB codes 1EJS and 1EJT for the Hisα219Asn and Hisα219Gln mutant, respectively, Table 1) [30]. Figure 11 shows how the mutated Glnα219 residue in 1EJT donates an H-bond through its -NH2 group to the water molecule coordinated to Ni(1), contributing to the building of the tetrahedral cluster of solvent molecules found in the native enzyme (compare with 1FWJ in Figure 4). The Hisα320Ala, Hisα320Asn, and the Hisα320Gln mutants display only a small change in Km, but a ⬃104 /105-fold decrease in kcat with respect to the wild-type enzyme [30,40]. Furthermore, their activity–pH profiles do not show the Hisα320 dependent pKa ⫽ 6.5 observed for wild-type urease [39,40], but rather a shift of the pH optimum to ⬃6 [30]. The structures of the Hisα320Ala, Hisα320Asn, and Hisα320Glu mutants (PDB codes 1KRC, 1EJU, and 1EJV, respectively, Table 1) have been determined [25,30]. In the case of 1KRC the Ni-bridging hydroxide is still found in place, while the major difference with the wild-type active site consists in the loss of the solvent molecules terminally bound to the nickel ions. This could be ascribed to the removal, in the mutant, of a residue capable to provide support for these molecules through the H-bonding network found in the wild-type enzyme, of which Hisα320 is a key player. In the case of 1EJU and 1EJV, the substitution of Hisα320 with Asn or Gln residues guarantees similar H-bonding properties compared with the native residue. Consistently, the active site structures reveal the presence of the four solvent-derived molecules in locations similar to those found in the wild-type (Figure 11, 1EJU). However, the active site flap becomes disordered and is not visible in the electron density, thereby hindering the localization of the Asnα320 or Glnα320 residues. In these mutants the capability of acid/base catalysis provided by the imidazole ring of Hisα320 in the native protein is compromised by the presence of the amide functional groups of the mutated residues, thereby justifying the drastic decrease of enzymatic activity. Met. Ions Life Sci. 2, 241–278 (2007)

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In the structure of KAU, the carboxylic group of Aspα221 is at H-bonding distance (3.0 Å) from the Hisα320 Nδ atom. This suggests that the latter is protonated, and therefore that the corresponding Hisα320 Nε atom is deprotonated at the crystallization pH of 7.5, very close to the optimum pH for catalytic activity (pH ⫽ 8). This could induce a role of proton acceptor for the Hisα320 residue during the catalytic cycle. Mutation of Aspα221 into alanine provokes a 103 decrease of the catalytic rate and a shift of the pH optimum to ⬃5 [30]. The structure of the Aspα221Ala mutant (PDB code 1EJR, Table 1) reveals a large disorder of the active site flap that contains Hisα320, while all the rest of the active site residues are essentially unchanged. This observation supports a role of Aspα221 in regulating the enzymatic activity through the modulation of the structural and reactivity features of Hisα320. Cysα319 is located on the flexible flap covering the active site of KAU. This residue is largely conserved in all urease enzymes, except for the enzyme from Staphylococcus xylosus, which has a threonine in this position [63]. Chemical modification of Cysα319 blocks enzyme activity [55,56], indicating that this residue is somehow involved in catalysis. However, the Cysα319Ala mutant is still ⬃50 % as active as the wild-type urease [57]. Structures of this mutant were determined at pH 6.5, 7.5, 8.5, and 9.4 (PDB codes 1FWB, 1FWA, 1FWC, and 1FWD in that order, Table 1) [26]. There were no significant differences among all these structures. The most evident differences between the structure of Cysα319Ala mutant and that of the wild-type enzyme involve a much-reduced mobility of the flexible flap covering the active site, still found in a ‘closed’ conformation, but displaying significantly reduced mobility. In contrast, the structures of Cysα319Asp (PDB code 1FWF), Cysα319Ser (PDB code 1FWG), and Cysα319Tyr (PDB code 1FWH) (see Table 1), which display, 0.03, 4.5 and 0%, respectively, of the activity observed for the wild-type, indicate a much higher mobility of the flap, but the same active site environment [26]. In the case of Cysα319Tyr, the flap is in the open conformation as for native BPU, an observation that was related to the presence of the bulky tyrosine side chain [26].

3.6. Structure of a Metal-Substituted Urease Since the discovery that urease, a hydrolytic enzyme, contains Ni2⫹ as the essential cofactor [12], the question about the role of this metal ion in the catalytic mechanism has intrigued the bioinorganic chemistry community. An approach to the solution of this problem has involved attempts to substitute the essential element with other ions such as Zn2⫹, Co2⫹, and Mn2⫹. Removal of both Ni2⫹ ions by treating JBU with EDTA at low pH causes irreversible denaturation of the protein [64]. Removal of a single Ni2⫹ ion by dialysis in the presence of citrate was reported [65], indicating that the two metal ions are bound with different affinity to the protein matrix. This result can now be interpreted on the basis of the crystal structures of KAU, BPU and HPU, which show that Ni(1) is bound to three protein residues (two histidines and one O-atom of the carbamylated lysine), while Ni(2) is additionally Met. Ions Life Sci. 2, 241–278 (2007)


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coordinated by the carboxylate group of an aspartate residue, therefore inducing a higher binding constant for the latter ion. The more labile metal ion in JBU can be substituted with Zn2⫹ or Co2⫹, using long-term dialysis [65]. The mixed-metal derivatives, as well as the derivative containing a single Ni2⫹ ion, are catalytically inert, indicating the essentiality of the presence of two Ni2⫹ ions in the active site for an efficient reaction mechanism. In 1995, just before the publication of the first crystal structure of KAU that revealed the presence of a carbamylated lysine as a Ni-bridging ligand, the requirement of CO2 in the form of a carbonate/bicarbonate buffer was reported to be essential during reconstitution experiments of apo-urease with Ni2⫹ ions [66]. However, only ⬃15% of the protein reconstituted in vitro is fully active and probably contains the carbamylated lysine and nickel ions as found in the crystal structures of the native urease proteins. The remaining ⬃85% of the reconstituted protein is present as an inactive form. This type of behavior also applies to urease reconstituted with Zn2⫹, Co2⫹, Mn2⫹, and Cu2⫹ [67], all of them possessing essentially no activity, with the exception of the Mn-derivative, which displays ⬃0.3% of the activity found in the native enzyme. The carbonate/bicarbonate in the reconstitution buffer can be substituted with CS2, to yield an inactive dithiocarbamylated Ni-bridging lysine derivative, and with vanadate (VO3⫺ 4 ), to yield an active form of the enzyme, possibly containing the vanadylated lysine derivative [68]. On the other hand, SO2, borate, sulfate, phosphate or molybdate had essentially no effect on the activation of apo-KAU. In the absence of CO2 in the reconstitution buffer, apo-urease binds Ni2⫹ to yield a distinct inactive Ni-containing species [66]. This latter species contains two Ni atoms, each coordinated by two histidines and 3–4 N/O ligands, as established by X-ray absorption spectroscopy [29], suggesting a similar active site as found in the structure of the native active enzyme, with the difference of having the carbamylated lysine oxygen atoms replaced by solvent molecules. The crystal structure of the inactive form of the Mn derivative of KAU was determined [29] (PDB code 1EF2, Table 1 and Figure 11). The structure features a similar arrangement of the active site as found in the native Ni-bound form, with some differences observed in the arrangement of the solvent molecules around the metal ions (shifted to much longer distances, 2.2–2.8 Å compared with 2.0–2.2 Å in native KAU) and a significant disorder of the Mn(2)-bound aspartate carboxylate group. The bottom line of these studies is that, in addition to the identity of the metal ions, the precise position of the protein ligands and solvent molecules around them is very important to achieve urease activity.

4.

THE STRUCTURE-BASED MECHANISM OF UREASE

It is generally accepted that, in the transition state of substrate hydrolysis, the planar urea molecule is converted to the tetrahedral (NH2)2C(O)(OH) ⫺ ion, Met. Ions Life Sci. 2, 241–278 (2007)

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CIURLI NH(Lys)

NH(Lys)

(His)N

N(His)

(His)N

Ni(1) H O

Ni(2)

OH2

H2O

Substrate binding

(His)N (His)N

N(His) O O(Asp)

N(His) Ni(1) H O

Ni(2)

N(His)

O

3 H2O C

H2 O

O(Asp)

NH2

C

NH2

H2 N

H2N

NH3 H2N-COOH

O

O

O

O

Alaα366

Flap closure

Flap opening

O

4 H2O NH(Lys)

NH(Lys)

O

O Ni(1) (His)N

Ni(2) O

O C NH3 N HN

O

O

(His)N

NH2

N(His)

(His)N

N(His)

(His)N

O(Asp)

O

Proton transfer

N(His) Ni(1) H O O C NH2

Ni(2)

NH2

N(His) O(Asp)

O

Alaα366

Alaα366

Figure 12. Structure-based urease catalytic mechanism of the enzymatic hydrolysis of urea.

having a central carbon and both N atoms formally hybridized sp3. The availability of the high-resolution structures of urease in the native state (PDB codes 1FWJ and 2UBP for KAU and BPU) and complexed with a substrate analog (BPU complexed with boric acid, PDB code 1S3T), with a tetrahedral transition state analog generated in situ by hydrolysis of the slow-reacting substrate phenylphosphorodiamidate (BPU complexed with diamidophosphate, PDB code 3UBP) and with a tetrahedral inhibitor (BPU complexed with phosphate, PDB code 1IE7) allows us to discuss the mechanism of catalysis on a solid structural basis (see Figure 12).

4.1.

Mechanism Step I: Binding of Urea to the Active Site

Theoretical calculations suggest that urea must first bind to the enzyme active site with the flap in the open conformation [69]. In this model, the urea-O atom binds to the more electrophilic and unsaturated Ni(1), with the consequent displacement of the two active site water molecules W1 and W3. The Ni(1)-bound urea O atom

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would be stabilized in this position through the formation of an H-bond donated by the NεH group of Hisα222, analogously to what is observed in the structures of native urease as well as BPU complexed with boric acid, diaminophosphate, phosphate, and acetohydroxamic acid. In this configuration, the urea NH2 group could form H-bonds with the Ni-bridging hydroxide WB and with the Ni(2)-bound W2 water, as suggested by recent calculations [70] that however did not take into account the presence of the conserved Alaα170 residue (the BPU sequence numbering will be used from now on). The carbonyl O atom of the latter could indeed very well act as an H-bond acceptor, as observed in the structures of BPU complexed with boric acid, diamidophosphate, and phosphate. The closure of the active site flap, which decreases the active site volume available for the substrate molecule, was found to facilitate the further binding of the urea NH2 group to Ni(2) to yield a urea molecule bridging the two Ni atoms [69], although this step could occur also in the absence of flap closure [70]. Closure of the flap would also be responsible for the stabilization of the catalytic transition state through the formation of multiple H-bonds with active site residues. The binding of the urea N to Ni(2) would be favored by the change of this N atom from ‘pseudo’ sp2 (with some sp3 character, as established by DFT calculations [69,70]) to pure sp3. The consequent loss of resonance delocalization energy of urea (⬃30–40 Kcal mol⫺1) would be compensated by the formation of the urea N–Ni(2) coordination bond. The hybridization change of the urea N (and possibly the formation of the urea N–Ni(2) bond) should directly follow, or be concomitant with, the nucleophilic attack on the urea C by an incoming hydroxide. It is also possible that the partial sp3 character of the N in unbound urea is enough to induce the coordination bond with Ni(2), thereby favoring nucleophilic attack on the urea C by loss of resonance energy and decrease of electron density on the C atom itself. The fact that B(OH)3 is a competitive inhibitor of urease, the isoelectronic structure of B(OH)3 and urea, the bridging-chelating binding mode of boric acid, the topology of the H-bonding network, the replacement of the labile water molecules in the active site by a neutral trigonal molecule, all provide evidence that gives support to the proposal of a mechanism involving a topologically similar binding of the substrate urea to the Ni ions in the active site. This substrate-binding mode involves a direct role of both Ni ions in binding and activating the substrate, therefore explaining the low reactivity of urease containing a single Ni ion. The position and orientation of the substrate are induced by the steric constraints as well as the asymmetric structural features of the active site, poised to donate H-bonds in the vicinity of Ni(1) and receive H-bonds in the vicinity of Ni(2). Urea is a poor chelating ligand because of the low Lewis base character of its NH2 groups; however, the formation of strong H-bonds with the nearby carbonyl oxygens could enhance the basicity of the NH2 group and facilitate the interaction of the amide nitrogen with Ni(2).

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4.2. Mechanism Step II: Nucleophilic Attack by the Bridging Hydroxide to Give the Tetrahedral Transition State The presence of the nonsubstituted Ni-bridging hydroxide in the complex of urease with B(OH)3 suggests a role for the Ni-bridging hydroxide WB as the nucleophile attacking the urea C atom to yield a tetrahedral transition state and intermediate. Such tetrahedral moiety could very well be represented by the structure of DAP bound to BPU. This scheme, first proposed in 1999 [31], is able to resolve several biochemical incongruities of the previously proposed mechanism, which involved as the putative initial step the action of a Ni(2)-bound terminal hydroxide reacting with a substrate molecule bound to Ni(1) through the carbonyl oxygen [14,18]. This latter proposal requires that the solvent molecule W2 terminally bound to Ni(2) is indeed the nucleophilic hydroxide ion, while all chemical evidence indicates that the pKa for such water is ⱖ10 [49] and therefore it must be protonated and neutral at the pH of maximal enzyme activity (pH ⯝ 8). On the other hand, the pKa for a nickel-bridging water molecule decreases significantly to very acidic values, while the pKa for the deprotonation of a metal-bridging hydroxide moiety is slightly lower than the pKa of the first ionization of a single ion bound to water [49]. Therefore, the estimated pKa for the deprotonation of the Ni-bridging solvent molecule (⬃9–10) guarantees that this moiety is already a hydroxide at the optimum pH for enzyme activity. The kinetic inertia of a doubly coordinated nucleophile could be easily overcome by the weakening of the Ni(1)–WB bond upon substrate binding, as suggested by kinetic data on the inhibition of KAU with fluoride [71], by the structure and reactivity of inorganic models [72], and by theoretical studies [69]. The high reactivity of the nickel-bridging hydroxide is also supported by the ability of the enzyme to hydrolyze PPD [53] and perhaps phosphate, where the enzyme would simply induce an oxygen atom (or hydroxide) exchange on the phosphate moiety. In this hypothetical framework, only molecules able to react with the bridging hydroxide may bind the enzyme in a tridentate mode, as observed for DAP and PHO. This would explain why DAP strongly inhibits urease when DAP itself is formed by enzymatic hydrolysis of PPD, (C6H5)–O–P(O)–(NH2)2, whereas it is a weak inhibitor if externally added to the native enzyme [53]. The lack of reactivity of B(OH)3 with the bridging hydroxide (placed at 2.1 Å from the B atom, in a direction almost perpendicular to the plane of the molecule) could be due to unfavorable symmetry and energy of the highest energy orbital carrying the two electrons necessary for the bond formation (the HOMO) on the nucleophile and the lowest energy empty orbital (the LUMO) on the inhibitor. The nucleophilic attack by the Ni-bridging hydroxide onto the sp2 carbon atom of urea yields a tetrahedral transition state containing an sp3 carbon. The structures of BPU complexes with diamidophosphate and phosphate well reproduce this state of the enzyme. In both structures the four oxygen or nitrogen atoms of the inhibitor lie in positions close to those of the four tetrahedrally arranged water Met. Ions Life Sci. 2, 241–278 (2007)


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molecules in the native enzyme [31]. The comparison of the DAP- and PHO-BPU structures also reveals why phosphate [52,62] binds much less strongly than DAP [53]. In the DAP complex the Ni(2)-bound DAP N atom donates two H-bonds to the carbonyl oxygen atoms of Alaα170 and Alaα366, while the distal DAP N atom is involved in H-bonding interactions with Ala α366 O, Glyα280 O and Hisα323 Nε [31]. The corresponding distal and Ni(2)-bound protonated phosphate oxygen atoms in the PHO-BPU complex can each only donate a single hydrogen, and the binding is hence weakened by an energy roughly equivalent to two or three H-bonds. Indeed, this optimization of H-bond donation from transition state to enzyme confers the observed specificity of urease: the substrate must be correctly oriented in the active site so to achieve the appropriate H-bonding network.

4.3. Mechanism Step III: Protonation of the Distal Urea Nitrogen Followed by Release of the First Ammonium Ion The nucleophilic attack onto the Ni-bridging urea molecule profoundly modifies the electronic structure of the substrate, and in particular, an increase of the pKa of the distal urea N atom, not involved in Ni-binding, is expected to occur. The consequent proton addition is certainly the driving force for the urea C–N bond breakage. Consistent with these considerations, DFT calculations have established that, upon nucleophilic attack of the bridging hydroxide on the urea C atom, the urea N atoms become sp3 and their negative charge density increases, favoring protonation [69]. The same calculations also established that protonation of the intermediate formed upon nucleophilic attack by the hydroxide on the substrate urea causes the breakage of the C–NH⫹3 bond and release of ammonia [69]. This step could be facilitated through the interaction of the nascent C–NH⫹3 group with Hisα323 Nε [31]. In this way, Hisα323 Nε would act as a base, stabilizing the proton on the distal urea N. The 103-fold reduction in kcat on mutation of Hisα320 in KAU (corresponding to Hisα323 in BPU) [30] could be explained in this context. The structure of the PHO-BPU complex also reveals that another active site residue, Alaα366, could be important in regulating the protonation state of the distal –NH2 group. This residue can adopt two different conformations (see models 2UBP and 3UBP in Figures 5 and 7) both thermodynamically stable, possibly constituting a sort of ‘molecular switch’ able to provide stabilization for the protonation of the distal urea NH2 group. A remaining point to discuss in the mechanism is the source of the proton that gives rise to the distal C–NH⫹3 group. On the basis of kinetic studies on KAU mutants it was proposed that Hisα323 is the proton donor to the urea–NH2 group [30]. This proposal requires a so-called reverse protonation hypothesis [14] in order to circumvent the problem of having a residue (Hisα323) reported to have a pKa ⫽ 6.5 in KAU [39, 40]) in a protonated state at the optimum pH for enzyme activity (⬃8) in order to deliver the proton to a urea-NH2 group and form ammonia. The Met. Ions Life Sci. 2, 241–278 (2007)

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result of such reverse protonation is that only about 0.3% of the enzyme molecules are active at pH 8. On the other hand, the PHO–BPU complex indicates that Hisα323 must be deprotonated at the crystallization pH of 6.3, and even more so at the optimal pH. In fact, the structural data for the PHO–BPU complex clearly reveal that the active site flap is open, and that the side chain of Hisα323 (even though not modeled because of disorder) must be located far from the active site, not able to interact with the phosphate moiety. If this residue were protonated at this pH, it could further stabilize the inhibitor binding through the formation of an additional H-bond with the distal phosphate oxygen. The fact that such stabilization does not occur is an indication that the pKa of Hisα323 may indeed be low, rendering it less likely to act as a proton donor during catalysis. In DAP-inhibited BPU, Hisα323 Nε is involved in an H-bond with the distal N atom of the Ni-bound DAP [31]. This interaction does not require the protonation of Hisα323 Nε, which could act as an H-bond acceptor from the distal -NH2 group of DAP [31]. In the structure of DAPinhibited BPU the active site flap is in the ‘closed’ conformation, allowing Hisα323 to approach the Ni environment in the active site. In this view, the combined action of flap closure and Hisα323 H-bonding acceptor capability appears to play a role of a perfectly tuned device able to select the access and stabilize the binding of molecules featuring, at the ‘distal’ position, at least two or better three hydrogen atoms, as in DAP and urea. On the basis of the structure of the DAP-BPU complex, it was proposed that the proton needed for protonation of the urea-NH2 group could come from the bridging hydroxide itself [31]. The higher pKa observed in the pH–urease activity profile (⬃9.5) corresponds to the estimated pKa for the bridging hydroxide, suggesting that indeed the bridging hydroxide moiety must carry a proton for the enzyme to work properly. The apparent difficulty to remove the H-atom from the bridging hydroxide of the catalytic intermediate, sometimes erroneously supposed to have a high pKa, can be disputed by considering that the proton would be released from a species containing an OH moiety bridging the two divalent Ni ions through the oxygen atom while being bound to an sp3 carbon. Such a moiety, essentially an alcohol, is expected indeed to be very acidic and prone to release the proton. One example of such a decrease of the pKa of metal-bound alcohols is provided by alcohol dehydrogenase, where the substrate alcohol experiences a large shift of pKa (from 16 to 6.4) upon binding to the active site Zn2⫹ ion [73]. The proton released by the bridging OH moiety could be transferred to the distal -NH2 group through the Oδ2 atom of Aspα363, a residue bound to Ni(2) through Oδ1 [31]. This proposal is suggested by the position of Aspα363 Oδ2, placed between the Ni-bridging DAP oxygen (at 2.5 Å) and the ‘distal’ DAP oxygen (at 3.5 Å), and is further supported by the observed conformational flexibility of the Cβ-Cα bond of Aspα363 in the structure of the BPU–AHA complex [33]. Recent calculations have shown that Aspα363 Oδ2 is deprotonated in the catalytically favorable binding mode of urea [70], thereby providing support for this group acting as a proton shuttle between the bridging -OH and the distal -NH2 group of the catalytic intermediate. Met. Ions Life Sci. 2, 241–278 (2007)


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CONCLUSIONS

Since the fi rst chemical synthesis of urea from ammonium cyanate in 1828 by Wöhler [74], since the fi rst isolation of a ureolytic bacterium, Micrococcus ureae, in 1837 by Schwann [75], since the fi rst crystallization of jack bean urease in 1926 by Sumner [11], which provided clear evidence that enzymes were actually proteins, and since the fi rst discovery by Zerner and coworkers in 1975 that nickel, a metal previously considered only for its toxicity, was indeed an essential cofactor of urease [12], research in this field has picked up at a steadily increasing pace, through a long and winding road of frustration and success. Much progress has been made in recent years towards a full understanding of the catalytic hydrolysis of urea, a fundamental process for both basic biochemistry as well as applied environmental, agricultural, and medical sciences, through the use of high-resolution protein crystallography. The results described in this review, as well as the detailed discussion of their possible implications, should be profitable for a wider and deeper understanding of the catalytic mechanism of this important enzyme, as well as providing more general ideas about the functioning of binuclear metallic hydrolases. These hypotheses should be useful for the development of drugs able to diminish the negative effects of urease in human and animal health [4,5,76] as well as in the agricultural environment [4,77,78].

ACKNOWLEDGMENTS This review is a result of the longstanding collaboration between the author and Dr Stefano Benini (now at York Structural Biology Laboratory, UK), and Professors Stefano Mangani (University of Siena, Italy), Wojciech R. Rypniewski (Polish Academy of Science, Poznan, Poland), and Keith S. Wilson (York Structural Biology Laboratory, UK). Many of the thoughts it contains are the products of strong and close interactions with these fine scientists as well as with all the author’s coworkers in Bologna, in particular Dr Francesco Musiani and Dr Barbara Zambelli. They are thanked for their work and inspiration. That said, responsibility for the opinions expressed here and whatever error this review contains rests entirely with the author.

ABBREVIATIONS AHA BME BPU

acetohydroxamic acid β -mercaptoethanol Bacillus pasteurii urease Met. Ions Life Sci. 2, 241–278 (2007)

274

DAP DFT EDTA EXAFS HPU JBU KAU MCD PDB PHO PPD RMSD UV

CIURLI

diamidophosphate density functional theory ethylendiamine-N,N,N⬘,N⬘-tetraacetate extended X-ray absorption fine structure Helicobacter pylori urease jack bean urease Klebsiella aerogenes urease magnetic circular dichroism protein data bank phosphate phenylphosphorodiamidate root-mean-square deviation ultraviolet

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7 Nickel Iron Hydrogenases Wolfgang Lubitz, Maurice van Gastel, and Wolfgang Gärtner Max-Planck-Institut für Bioanorganische Chemie, Striftstrasse 34-36, D-45470 Mülheim an der Ruhr, Germany

1. INTRODUCTION TO HYDROGENASES 2. BIOCHEMISTRY AND MOLECULAR BIOLOGY 2.1. Cellular Location, Structure, and Composition of [NiFe] Hydrogenases 2.2. Hydrogenase-Encoding Genes 2.3. Maturation Pathway 2.4. Genetic Manipulation 2.4.1. Electron Transport Pathway Mutations 2.4.2. Proton Transport Pathway Mutations 2.4.3. Mutations Affecting Oxygen Tolerance 2.4.4. Mutations Directed Toward the Active Site 3. CRYSTALLIZATION AND X-RAY STRUCTURE ANALYSIS 4. SPECTROSCOPIC INVESTIGATIONS 4.1. FTIR Spectroscopy 4.2. X-Ray Absorption Spectroscopy 4.3. EPR Spectroscopy 4.3.1. The Intermediate Ni-C State 4.3.2. The Oxidized States 4.3.3. Interaction with the Protein Environment 5. ELECTROCHEMISTRY 6. HYDROGENASE FUNCTION AND THE CATALYTIC CYCLE



280 283 283 284 285 287 289 289 290 290 291 295 295 299 300 303 305 306 306 309

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CONCLUSIONS AND OUTLOOK ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

312 314 314 315

INTRODUCTION TO HYDROGENASES

Nickel is an essential constituent of a large subgroup of hydrogen-metabolizing enzymes. These enzymes, hydrogenases, catalyze one of the simplest molecular reactions, the heterolytic conversion of hydrogen into protons and electrons and the reverse reaction, i.e., the generation of hydrogen: H2 H H 2 H 2 e Hydrogenases are widespread in bacteria, archaea, and eukarya. They can be classified according to the composition of the active site in [NiFe], [FeFe] or [Fe] hydrogenases. A subgroup of the first is formed by the [NiFeSe] hydrogenases, in which one of the cysteine residues that serves as a ligand to the two metal ions is exchanged to a selenocysteine. Another classification can be made according to the redox partners, which in most cases is either NAD(P) or a b- or c-type cytochrome. Hydrogenases are found in the periplasm or in the cytosol, either in soluble form or membrane-attached. In eukaryotic cells hydrogenases are often located in specialized cellular compartments [1,2]. The most important function of hydrogenases is probably to balance the redox potential within the cell, and also to provide energy by the dissociation of molecular hydrogen. However, the same principle to bind hydrogen in the active site as during the enzymatic process has also been utilized to sense the presence of hydrogen and regulate gene activity to produce hydrogenases. Sensing of hydrogen by the common [NiFe] active center then initiates a signaling process (via the widespread prokaryote two-component signal transduction pathway) which results in a physiological response of the organism [3]. Accordingly, hydrogenases are often classified as membrane-bound hydrogenases (MBH), soluble hydrogenases (SH) or regulatory hydrogenases (RH). The activity of hydrogenases can be measured by several assays which are described in detail in the literature [1,4]. The catalytic activity of [NiFe] hydrogenases has been found to be lower than that of [FeFe] hydrogenases [5]. This has, however, recently been questioned by Armstrong and coworkers who were able to show that in protein film voltammetry experiments high turnover rates are also obtained for [NiFe] hydrogenases [6]. Met. Ions Life Sci. 2, 279–322 (2007)

NICKEL IRON HYDROGENASES

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The strict requirement of anaerobic conditions to prepare functionally active enzymes, their low concentrations in living cells, and the complex maturation pathway might have been responsible for the difficulties to characterize the role of nickel for the enzymatic activity. The enzymatic function was first described in 1931 [7], whereas the first identification of nickel as a constitutive component in hydrogenases was reported in 1965, based on supplementation experiments of the growth media [8]. The involvement of nickel in the redox processes that take place during the conversion of hydrogen, has been established by 61Ni isotope labeling and EPR spectroscopy in the early 1980s by the work of Thauer and Albracht on the enzyme from a methanogenic bacterium [9,10]. Le Gall et al. reported the same effects for D. gigas hydrogenase [11] and Cammack and coworkers [12] carried out redox titrations at different pH values and also discussed the possibility of nickel being involved in the catalytic cycle. EPR has also been very useful to investigate the redox states of iron–sulfur clusters present in the enzyme [13,14]. These contributions demonstrate the effectiveness of EPR spectroscopy for the study of the electronic properties of this enzyme [13,15,16] in particular when combined with advanced techniques such as multifrequency, high-field pulsed EPR, and double resonances such as ENDOR [17]. Based on the data elucidated from spectroscopy and the functionality of the enzyme, a first model for the activation/inactivation and the catalytic cycle of [NiFe] hydrogenase was established in 1985 [18]. The first crystallographic structure of a [NiFe] hydrogenase was reported by Volbeda et al. in 1995 [19,20]. This work clearly showed that the active site contains a heterobinuclear [NiFe] center anchored in the protein via cysteine residues. Surprisingly, three small diatomic molecules were found as ligands to the iron that were later identified as two CN and one CO by FTIR [21,22]. A schematic overview of the protein subunits, active site and the electron/proton pathways and a gas channel for H2 is shown in Figure 1. In the oxidized, as-isolated form the enzyme is catalytically inactive and exists in a mixture of two states (Ni-A, unready and Ni-B, ready), which differ in activation time. Upon reduction the enzyme passes through the states Ni-SU, Ni-SIr, the one-electron reduced states belonging to Ni-A and Ni-B, respectively. At room temperature the Ni-SIr can undergo a structural change into another state called ‘silent active’, Ni-SIa. Upon further reduction of the enzyme in the Ni-SIa state, the one-electron reduced Ni-C and two-electrons reduced Ni-R states are reached. Whereas the Ni-SU, Ni-SI and Ni-R states are ‘EPR-silent’ in typical EPR experiments at X-band, the Ni-A, Ni-B and Ni-C states yield pronounced EPR signals characteristic for paramagnetic nickel states (electron spin S 1兾2). The Ni-C state is light sensitive and yields several Ni-L states, which are also paramagnetic (S 1兾2). A new field in hydrogenase research was opened with the application of FTIR spectroscopy [23]. The studies performed on different enzymes [20,24–26] showed bands in the 1800–2100 cm1 spectral region which were assigned to Met. Ions Life Sci. 2, 279–322 (2007)

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2H

H+ + Mg2

[NiFe] center

H+ CN S x Fe CO Ni S S CN H2

Proton channel

S

H2

H+ e– S Fe S Fe S Fe S

Fe

H2 Hydrogen channel

e– FeS

e– e–

FeS

+

H

Large subunit (~60kD)

Electron acceptor/ donor

Small subunit (~30kD)

Figure 1. Schematic view of the catalytic [NiFe] hydrogenase showing the two subunits, the active site [NiFe] cluster, the hydrogen channel and the proton transfer channel (with a Mg2 included) and the three iron–sulfur clusters in the small subunit that constitute the electron transport chain (to an exogenous electron acceptor/donor), see ref. 35.

vibrations of CN and CO ligands at the iron. These bands were found to shift significantly when the enzyme was reduced or oxidized either electrochemically or by gas exchange (H2, N2 or O2). The observed effects were later extensively used by several groups to characterize the intermediates of the enzyme’s activation/inactivation and catalytic cycle. Whereas EPR is sensitive only to the paramagnetic states (mostly S 1兾2 states) FTIR has the advantage of being sensitive to all redox states. Important contributions for a structural and functional understanding of the enzyme have also come from other spectroscopic techniques such as X-ray absorption [27] and Mössbauer spectroscopy [28]. The conversion of ortho to para hydrogen has been investigated [29] and experiments using mass spectroscopy have recently been reviewed [30]. Furthermore, the development of new electrochemical methods by Armstrong et al. helped to acquire a better understanding of the enzyme’s redox chemistry and its possible electrocatalytic applications [31–33]. The crystal structure, together with functional spectroscopic and electrochemical investigations as well as biochemical and genetic studies, laid the basis for our Met. Ions Life Sci. 2, 279–322 (2007)


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present knowledge about the [NiFe] hydrogenases. These aspects will be described in this article with special emphasis on results obtained during the last decade. Some of the work has already been reviewed and the reader is referred for the older literature and more extensive information to the two books edited by Lancaster [34] and by Cammack et al. [35] and to several review articles [1,2,36–43].

2.

BIOCHEMISTRY AND MOLECULAR BIOLOGY

2.1. Cellular Location, Structure, and Composition of [NiFe] Hydrogenases The majority of [NiFe] hydrogenases are built as heterodimers, consisting of a large and a small subunit of ⬃45–70 kDa and ⬃30–35 kDa, respectively. An analysis of the protein sequence, the folding pattern and the three-dimensional structure revealed similarities between the [NiFe] hydrogenases and other energy-providing, redox-active protein complexes such as NADH:ubiquinone oxidoreductases, except for the architecture of the active site [44–48]. In fact, the structure and also biosynthesis of the small subunit containing the [FeS] clusters follows a general blueprint for this type of proteins. Hydrogenases from bacteria are found in the cytosol and also in the periplasm. The periplasmic hydrogenases (in soluble or membrane-attached form) carry signal peptides at the N-terminus of the small subunit, allowing the transport through the inner membrane by the TAT (twin-arginine transport, recognition motif RRxFxK) pathway [2,49], whereas hydrogenases located in the cytoplasm are void of these signal sequences. In cases where hydrogenases are located in special cell compartments (chloroplasts or the so-called hydrogenosomes), similar signal peptides or recognition motifs can be identified that ensure correct transport [50,51]. The small subunit holds three iron–sulfur clusters of various compositions (as an example, the most intensely studied hydrogenases from the Desulfovibrio species carry a proximal and a distal [4Fe4S] cluster, flanking a central [3Fe4S] cluster). This arrangement, and a putative binding site for a redox-active compound, most probably serves as an electron transport pathway (see also below). The most common redox partners are cytochrome b, cytochrome-c3 or NAD. The active site, in the large subunit and close to the interface with the small subunit, holds a nickel and an iron atom in a distance which varies between ⬃2.5 and ⬃3.0 Å depending on the redox state of the enzyme. The electronic configuration of the binuclear complex is modulated by three ligands, two CN and one CO group, attached to the iron atom. These substituents are excellent probes for infrared-spectroscopic identification of the various oxidation states of the hydrogenase [21] (see also Section 4.1). Besides the nickel and the iron ion, at least one additional metal ion, magnesium, has been identified from the crystal structure analysis. It Met. Ions Life Sci. 2, 279–322 (2007)

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has been proposed that the Mg2 ion is part of the proton-translocating channel to/from the active site (see also Figure 1) [36,39]. The above mentioned dimeric structure of [NiFe] hydrogenases is found for many of the periplasmic and also the cytosolic species, soluble or membrane attached and irrespective of their function to metabolize hydrogen or only to serve as a hydrogen sensor (see below). Besides this widespread structural motif, several hydrogenases are known to be composed as multi-protein complexes. The hydrogen sensing enzymes play an important role in hydrogen metabolism. Due to the extremely complex and energy-consuming biosynthesis of hydrogenases (see below), a rigorous control on the transcription of genes encoding the hydrogenase subunits and the maturating proteins is necessary. This function is fulfilled by hydrogen-sensing proteins that do not convert hydrogen, but initiate upon its detection transcriptional activity, accordingly being named regulatory hydrogenases. These proteins make use of the same subunit architecture as the catalytic proteins, also containing a [NiFe] active site, but upon binding of hydrogen interact with a protein, that carries a histidine kinase activity. Histidine kinases and the directly interacting partner proteins, their cognate response regulators, are highly conserved in their structure and function within prokaryotes, and are widespread to detect a broad variety of external stimuli [52]. In case of the hydrogen detection mechanism, the histidine kinase changes its phosphorylation state (becoming either phosphorylated or dephosphorylated at a fully conserved histidine), which in turn allows interaction with the response regulator protein. During this protein–protein interaction, the phosphate group is transferred to the response regulator, which then, in its activated state, can interact with hydrogenase-gene-specific promoters to initiate gene transcription and hydrogenase biosynthesis [3]. Recent experiments give evidence that the sensory function is accomplished by a high-molecular-weight complex of up to 350 kDa which contains a number of additional proteins involved in this regulatory process [53].

2.2.

Hydrogenase-Encoding Genes

A wealth of information on the sequence and arrangement of genes encoding for structural and accessory proteins has been obtained in recent years, based on the ever-growing knowledge from sequencing of entire genomes. The most recent, most comprehensive compilation of sequence information [1] from 2001 lists more than 100 different hydrogenase genes, and since then more than 30 new entries can be found in the various databases (e.g., http://www.ncbi.nih.gov/; http://www.expasy.org/sprot/). In prokaryotes, genes encoding the small and the large subunit of [NiFe] hydrogenases are arranged in operons, often followed by various numbers of genes encoding maturation proteins and the regulatory – hydrogen sensing – proteins. Met. Ions Life Sci. 2, 279–322 (2007)

NICKEL IRON HYDROGENASES Hydrogenase 1 (hya)

285

A SSU

B LSU

D Prot. 1000 bp

Hydrogenase 2 (hyb)

O SSU

C D FG LSU Prot. ~5kbp

E

D C B A

Accessory Proteins (hyp)

E LSU Hydrogenase 3 (hyc)

G

I

F

SSU Prot. Accessory Protein (hyp)F

Figure 2. Organization of the structural genes of hydrogenase 1 (hya), 2 (hyb), and 3 (hyc), and the genes for the accessory proteins (hyp) in E. coli. Prot: protease. Genes for homologous proteins are indicated by the same greyscale. Adapted from [178] by permission of Elsevier GmbH/Spektrum Akademischer Verlag, Heidelberg.

In some organisms, the structural genes and those encoding the hydrogenmetabolizing machinery are even found organized on plasmids. For the various genes encoding different types of proteins (structural, accessory, regulatory proteins), separate promoters have been identified. This clustered organization has facilitated in many cases the search and identification of hydrogenase-related genes. Best characterized prokaryotes are E. coli [54], the various Desulfovibrio species [40,55] and Thiocapsa roseopersicina [56]. A schematic overview of the organization of the structural and auxiliary genes in E. coli is given in Figure 2.

2.3.

Maturation Pathway

The introduction of nickel and iron into the enzyme and the generation of the unusual substituents at the iron atom (carbon monoxide and the toxic cyanide group), together with the transport to the inner membrane or the periplasmic space proceeds via sophisticated and complex biosynthetic pathways. Pioneering work has been performed by the group of Böck on the hydrogenases of E. coli [54], followed by the studies of other groups on various Desulfovibrio species [57] and on Ralstonia eutropha [55]. Genetic analysis in E. coli revealed the presence of three hydrogenases (labeled as 1, 2, and 3). The study of the maturation proteins yielded a gene cluster hypA–E, and an additional gene, hypF, which were found to be essential for the biosynthesis of all three [NiFe]-hydogenases present in E. coli (hyp, hydrogenase pleiotropic). Detailed analysis showed an isoenzyme-specific function for hypA and hypC, causing an immature hydrogenase 3 (see Figure 2) upon selective deletion of these genes, whereas deletions in the other genes yielded a pleiotropic effect to all three hydrogenases. Interestingly, homologous genes for hypA and hypC were found for the maturation of hydrogenase 2, namely hybF Met. Ions Life Sci. 2, 279–322 (2007)

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and hybG, explaining the observation that the deletion of hypA and hypC causes defects in the biosynthesis of hydrogenase 3, but not of hydrogenase 2. The above-mentioned toxicity of the iron-attached carbon monoxide and cyanide substituents remained an enigma in the biosynthesis of hydrogenases until the finding of sequence similarities between HypF and O-carbamoyltransferases. Carbamoyl phosphate had been proposed as the precursor of these ligands, since a knock-out of the carbamoyl phosphate synthase impairs not only pyrimidine synthesis, but also that of hydrogenases [58]. Continuing investigations, however, raised doubts to the same origin of both substituents [59]. As an additional problem, if different synthetic routes for the two types of ligands are proposed, the unequivocal stoichiometry of 2:1 has to be ensured. For the process of ligand incorporation, the formation of a complex between HypE and HypF takes place. It could furthermore be shown that the assembly of the active site, i.e., first the incorporation of iron and its furnishing with the three diatomic substituents, followed by the incorporation of nickel is a prerequisite for the correct ongoing processing of the large subunit. Still, the involvement of a scaffolding protein in this process is discussed. The incorporation of iron precedes that of nickel, as can be seen from nickel-deficient cells, which still hold the iron incorporated into the large subunit. However, no intermediates in this assembly process have yet been isolated in sufficient amounts for a detailed biochemical or spectroscopic analysis. Both HypA and HypB are involved in the incorporation of nickel into the active site. However, hypB-deletion mutants can still synthesize functional hydrogenases under conditions of high nickel concentrations in the medium. One function of HypB is apparently to serve as a nickel-storage protein, the other – probably in a complex with HypA – is the incorporation of nickel. The uptake of nickel ions into cells is well studied and is outlined in a number of concise reviews, e.g., [60,61]. In E. coli, the ABC-transporter pathway is used for the reception of Ni (the same system also shows binding capacity for Co, Cu, and Fe, however with ⬃ 10-fold lower affinity). The transport system consists of a five-protein complex NikA-E that binds Ni in the periplasm (NikA) and transports it via formation of a pore (NikB and NikC) into the cytosol where it is further transported under ATP hydrolysis (NikD and NikE) to the various Ni requiring proteins. The second relevant Ni uptake process utilizes permeases, intrinsic pore-forming membrane proteins, e.g., HoxN in Ralstonia eutropha. This protein has been reported to be specific for Ni, and does not bind Co [62–64]. Only after completion of the binuclear metal center, the large subunit is cleaved at the C-terminus by a specific protease (e.g., HycI for hydrogenase 2 and HybD for hydrogenase 3 in E. coli [65,66]) at a position directly after one of two cysteine residues that are part of the active site ligands. For the cleavage reaction, the protease HybD forms a complex with HybC, which serves as a chaperone. After cleavage of the C-terminal extension, the large subunit forms a complex with the independently synthesized small subunit. The formation of the Met. Ions Life Sci. 2, 279–322 (2007)


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three [FeS] clusters in the small subunit is accomplished by a general pathway for this type of cofactors. Three systems are described for prokaryotes (discussed in [67]). Interestingly, in eukaryotes, all [FeS] clusters are built in the mitochondria, irrespective of their final location in the cytosol or in the nucleus [68]. Following identification of the signal peptide at the small subunit, the completely assembled holo-enzyme is then transferred to the cytoplasmic membrane. After passing the membrane, the hydrogenase is fixed at the periplasmic side. Many [NiFe]-hydrogenases carry a hydrophobic peptide at the N-terminus of the large subunit that serves as an anchor to hold the hydrogenase at the periplasmic surface of the membrane. Additional fixation takes place by complex formation with a membrane-associated cytochrome c or a membrane-intrinsic cytochrome b, serving as the electron acceptor. A schematic overview on the maturation pathway is given in Figure 3.

2.4.

Genetic Manipulation

The complexity of hydrogenase maturation has turned out to be a major obstacle for genetic manipulation and for the routine production of large amounts of hydrogenases by inducible expression in homologous or heterologous hosts. Beginning in the early 1990s, knock-out and site-directed mutations were performed, mostly in E. coli, in various Desulfovibrio species [40] and in Ralstonia eutropha [55], as well as in Thiocapsa [56]. Initially generated deletion mutants were of only limited information, since at most the presence and the number of hydrogenases in a certain organism could be determined. Deletion of hydrogenases also yielded information on the hydrogen metabolism from headspace analysis, i.e., whether a certain hydrogenase functions in hydrogen uptake or release mode [49]. Yet, these deletion mutants can now serve as hosts in cases where site-directed mutants are being constructed, allowing the isolation of only the mutated protein without ‘contamination’ of the wild-type. Genetic studies in less well characterized organisms require the construction of appropriate plasmids to introduce foreign DNA into the bacterium. In the beginning, naturally occurring ‘broad host range’ plasmids were simply modified by the insertion of additional sequences allowing cloning in E. coli (generation of shuttle vectors). A family of better tailored vectors of lower size, which are based on a plasmid from D. desulfuricans, has recently been described [69]. For molecular biology experiments to be performed in Ralstonia (which mostly concentrate on the hydrogen sensing protein), the organization of structural and maturation genes in large operons has been employed. A mega-plasmid was constructed with all essential genes that allows easy access to the various genes for mutagenesis experiments [70]. The general method of introducing foreign DNA into these bacteria is still via conjugation that results in an autosomally replicating plasmid or in the stable integration of foreign DNA in the genome. Met. Ions Life Sci. 2, 279–322 (2007)

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Figure 3. Model of the maturation pathway for a typical, membrane-bound [NiFe] hydrogenase. Adapted from [179] by permission of Elsevier GmbH/Spektrum Akademischer Verlag, Heidelberg.

Only few examples have been reported where DNA insertion was accomplished by electroporation [69]. More advanced studies now make use of an additional modification of recombinant hydrogenases such that an affinity tag is added to the protein, allowing a much more convenient purification of the expressed protein. Still a major problem in mutagenesis studies is the fairly low concentration of the mutated hydrogenase in the currently investigated bacteria, and – in the case Met. Ions Life Sci. 2, 279–322 (2007)


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of several of the Desulfovibrio species studied – the relatively low cell density. No strong induction has been obtained up to now. An exception is D. fructosovorans. This bacterium has several advantages such as being relatively easily transformed and, like other organisms, synthesizes soluble hydrogenases. Furthermore, its capability to utilize fructose as energy source results in higher cell densities and also allows the study of hydrogenases that are partially impaired in their function due to an introduced mutation. Yet, with the successful generation of site-directed mutations in the various hydrogenase-containing organisms, several functional problems could be addressed. The most important questions, of course, are those directed toward the function of the active site itself and those regarding the transport pathways or channels for the different substrates and products of hydrogen conversion, i.e., the pathways for electrons and for protons, and a channel allowing diffusion of hydrogen to the active site (see Figure 1).

2.4.1.

Electron Transport Pathway Mutations

One of the earliest mutations performed was the conversion of the central [3Fe4S] cluster in the small subunit into a [4Fe4S] cluster (see Figure 4 in Section 3). A comparison of the protein environment around that iron–sulfur cluster revealed the presence of a proline residue at a position where [4Fe4S]-binding proteins carry a cysteine, serving as additional ligand. It should be kept in mind that structural assignments of single amino acids to be subjected to mutagenesis relies on protein structures from D. gigas [71], D. vulgaris Miyazaki F [72] or D. fructosovorans [73], requiring a sequence alignment with the hydrogenase from, e.g., R. eutropha, if the mutations are planned to be performed in this protein. In fact, conversion of the identified proline into a cysteine in Desulfovibrio resulted in the formation of a central [4Fe4S] cluster [74]. This position was selected for mutagenesis since the [3Fe4S] cluster was shown to have a high midpoint potential. In fact, the mutation resulted in a lowering of the potential by ⬃300 mV, but did not lead to strong changes in the spectroscopic properties and altered the catalytic properties of the mutated enzyme only slightly.

2.4.2.

Proton Transport Pathway Mutations

Besides the gas channel, which has been investigated mostly by structural means, and the electron pathway via the iron–sulfur clusters, the third important transport pathway for reagents in hydrogenase activity is the transfer of protons between the active site and the surface. One may assume that, as in other proton transporting proteins, a pathway consisting of polar or charged amino acids exists, and in fact a polar network had been identified in the crystal structures of various hydrogenases, including a magnesium ion that holds in place several glutamate Met. Ions Life Sci. 2, 279–322 (2007)

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residues. A most promising candidate is Glu25 in D. fructosovorans which is assumed to be dissociated and is located relatively close to the active site. Exchanges were performed into aspartate (shorter side chain) and into glutamine, removing its dissociation capacity [75]. The E25Q mutant showed FTIR and EPR signals very similar to the wild-type protein, both in the oxidized and the reduced state, however, exhibited entirely different enzymatic properties. This mutant did not show any hydrogen uptake, or deuterium/proton exchange, which are indicative for a functional hydrogen transport. Investigations on the transport processes of electrons, protons, and hydrogen have intensified strongly in the very recent years thanks to more convenient cloning facilities. Accordingly, additional residues involved in proton transport have been identified in Ralstonia, the most fascinating one being R40 (having been converted into leucine [76,77]). This mutant is completely impaired in hydrogen uptake, but still shows the capacity to exchange protons with deuterons.

2.4.3.

Mutations Affecting Oxygen Tolerance

Also of particular interest is the oxygen tolerance of some hydrogenases, best studied for those from Ralstonia, in particular from the point of a possible technical application [78]. Since the active site is deeply buried within the protein, the hydrophobic channel (see Figure 4A in Section 3), which extends from the surface of the enzyme to the active site and which has been shown to be sufficiently large for the diffusion of hydrogen [73], was targeted by mutagenesis in the hydrogen sensing protein from Ralstonia [79]. This channel is probably also large enough for the passage of dioxygen, carbon monoxide, and nitrogen, which might cause the inactivation of hydrogenases upon exposure to air. The hydrogen-sensing protein from Ralstonia is insensitive toward oxygen exposure, and a comparison of its sequence to oxygen-sensitive hydrogenases, for which three-dimensional structures are also available, indicated two bulky residues in the proposed gas channel. A stepwise exchange into amino acids with smaller side chains (I62V and F110L) caused increasing oxygen sensitivity.

2.4.4.

Mutations Directed Toward the Active Site

Amino acids in the direct vicinity of the cyanide or carbon monoxide ligands were also addressed by mutagenesis in order to learn whether a hydrogen bonding network influences the electronic configuration of the active site (Val78, Pro498, and His499 in D. fructosovorans) [80], albeit the mutations caused relatively small effects on the catalytic activity of the modified hydrogenase. The directly adjacent cysteines in the active site, serving as ligands for the dinuclear metal center were mutated in the cytosolic (SH) hydrogenase of Ralstonia [76]. In all cases the assembly of these mutated proteins was impaired with deleterious effects on Met. Ions Life Sci. 2, 279–322 (2007)


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the enzymatic activity. An interesting mutation in direct vicinity of the binuclear metal center was performed again in the hydrogen-sensing protein from Ralstonia. This protein carries a glutamine residue at a position where catalytic hydrogenases show a histidine. An exchange of this glutamine into histidine or asparagine affected the stability of the mutated proteins, which still showed regulatory and enzymatic activity. An exchange into glutamate yielded an entirely unstable enzyme [81]. EPR measurements together with theoretical calculations gave an explanation of the effects of lower stability caused by the His and Asn mutations: a weak hydrogen bond is formed to one of the cysteines, which serve as ligands to the metal ions of the active site. All of the approaches employing site-directed mutagenesis are still at their beginning, and many more positions will and have to be addressed to understand the role of particular amino acids in greater detail. As outlined above, the relatively low yield of hydrogenase biosynthesis in their endogenous hosts, the complex maturation process, the oxygen sensitivity, and the difficult, time-consuming purification still represent major drawbacks for an efficient study of these exciting enzymes. Some of these obstacles (oxygen sensitivity, affinity-tag-based purification) have been addressed in recent years. Efforts have been performed to a heterologous expression that might facilitate the generation of larger amounts of recombinant hydrogenases, in their wild-type as also in mutated form. A recent approach performed for Ralstonia aimed at the construction of a mega-plasmid carrying all essential genes for hydrogenase generation and maturation [70]. Similarly, heterologous expression of Fe-only hydrogenases from Chlamydomonas reinhardtii and Senedesmus obliquus in Clostridium acetobutylicum have recently been reported [82], and can be assumed to be also suitable for the expression of [NiFe] hydrogenases. Further future perspectives in hydrogenase production were recently outlined [83,84].

3.

CRYSTALLIZATION AND X-RAY STRUCTURE ANALYSIS

The first crystallization of a [NiFe] hydrogenase was reported in 1987 [85], but an X-ray crystallographic structure did not appear before 1995, when Volbeda et al. published the structure of the enzyme from D. gigas [19,20]. Before this time little was known about the exact composition of the active site, except that it contained nickel [9,10]. Surprisingly, two metals were found, the other turned out to be iron. Since then, several [NiFe] hydrogenases from other organisms have been crystallized; an overview is given in Table 1. A compilation of the structural features of the [NiFe] hydrogenases is shown for the enzyme from D. gigas as an example in Figure 4. All enzymes have conserved coordination geometry of the active site. The nickel is coordinated by four cysteines, two of which form a bridge to the iron atom. In the class of [NiFeSe] hydrogenases one of the terminal cysteines is Met. Ions Life Sci. 2, 279–322 (2007)

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Table 1.

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Crystal structures of [NiFe] hydrogenases from different organisms.

Organism (functional state of the enzyme) D. gigas (Unready) D. gigas (Unready) D. gigas (Active) D. gigas (Unready) D. vulgaris MF (Ready) D. vulgaris MF (Active) D. vulgaris MF (Inhibition) D. vulgaris MF (Unready) (Ready) D. fructosovorans (Unready) D. fructosovorans (Ready) D. fructosovorans (Unready) D. baculatum (Active) a D. desulfuricans

Resolution (nm)

Year

PDB

Reference

MR MR

0.28 0.25 0.27 0.24 0.18 0.14 0.12–0.14 0.10–0.14

1995 1996 1998 2005 1997 1999 2002 2005

1FRV 2FRV 1CC1 1YQ9 1H2A 1H2R 1UBH-U 1WUI-L

[19] [20] [86] [71] [88] [87] [95] [72]

MR MR MR MR MR

0.27 0.21 0.18 0.22 0.18

1997 2005 2005 1999 2001

1FRF 1YRQ 1YQW 1CC1 1E3D

[73] [71] [71] [86] [89]

Method MIR MR MR MR MIR

a

[NiFeSe] hydrogenase. MIR multiple isomorphous replacement, MR molecular replacement. PDB (protein data bank file): http://www.rcsb.org

replaced by a seleno-cysteine [86]. A third bridging ligand X is present in the ready and unready oxidized states, which is absent in the X-ray structure when the enzyme is reduced [86,87]. This bridging ligand in the oxidized state has been postulated to be an oxygen [20] or a sulfur species [88,89]. Spectroscopic results using 17O labeling [90,91] support the former hypothesis for two of the enzymes, but the presence of a sulfur bridge in the inactive enzyme for other species cannot be completely excluded. Three out of the four sulfur atoms and the additional bridging ligand form the base plane of a square pyramid of the Ni coordination sphere, the fourth sulfur occupies the axial position. Furthermore, the crystal structure showed that the iron is carrying three diatomic ligands that were later identified by FTIR spectroscopy to be two CN and one CO [21]. In the oxidized (as isolated) state the iron is six-coordinated whereas the nickel is fivecoordinated. The structure of the active [NiFe] site is shown in Figure 4B. Whereas the [NiFe] center is located in the large subunit, three iron–sulfur centers (two [4Fe4S] flanking one central [3Fe4S]) are present in the small subunit (Fig. 4A,C). It has been concluded that the iron–sulfur centers constitute an electron transport chain extending from the interface of the two subunits close to the active site to the protein surface. In the structure another metal ion (Mg2) has been found that is probably involved in the transport of protons to the active site (Figure 4A,D). Met. Ions Life Sci. 2, 279–322 (2007)


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Figure 4. X-ray crystallographic structure of [NiFe] hydrogenase from D. gigas [19,20]. (A) Structure of the protein with subunits and domains, the NiFe center (green, red), the Mg (light blue) and the 3 FeS clusters (red/yellow). Hydrophobic channels are indicated in blue, which were probed by Xe atoms (blue spheres) introduced in the crystals. (B) Active [NiFe] cluster with ligand surrounding (D. gigas numbering); X is an oxygenic species in the oxidized state, in the reduced state X is missing in the structure. (C) Possible electron transfer path from the active site via the [FeS] centers to the protein surface (dotted line). (D) Possible proton transfer path from the active site via Mg2 and structural water molecules to the protein surface. Note that several glutamic acids (Glu18, 46, 321) and a His(536) are proposed to be involved in this process. Part of the gas channel is also shown. Reprinted with permission from [36] (A, C, D); Copyright John Wiley & Sons, Ltd, 2001.

Another interesting observation is the presence of gas channels, indicated in Figure 4A. One extends from the surface of the protein to the deeply buried active site. The identification of these channels was accomplished first by a cavity calculation on the structure of D. gigas hydrogenase with a probe radius of 0.1 nm. Additional information was obtained by X-ray analysis of single crystals which Met. Ions Life Sci. 2, 279–322 (2007)

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had been exposed to high pressure Xe gas [73], causing gas atoms to diffuse into the channels (Figure 4A). Strikingly, the channel seems to end at the vacant sixth coordination position of the nickel. It has been speculated that this is the channel used by hydrogen, but may also allow access of other gas molecules such as CO and O2 which lead to inhibition of the enzyme. Recently, new crystal structures became available for both the ready and the unready states [71,72]. Though the structure of the ready state was essentially a confirmation of the earlier structures that of the unready state held a surprise. The electron density for the third bridging ligand was found to be elongated, and it was suggested that this ligand is possibly hydroperoxide, OOH [71]. Furthermore, extra density was found in these structures [71,72] near the cysteine sulfur atoms, indicating the formation of sulfoxide species possibly resulting from trapped O atoms as recently discussed by Albracht, Armstrong, and coworkers [92–94]. It was proposed that the type of bridging ligand may be the cause for the slow activation of the unready state and the fast activation of the ready state [71]. Ogata et al. [72] compared their highly resolved Ni-A structure with that of the CO-inhibited enzyme [95] and could show that the extra density is in the same position as that of the CO ligand that blocks the sixth coordination position of the nickel (see Figure 5). This would prevent binding of dihydrogen.

Figure 5. Comparison of the X-ray crystallographic structures of Ni-A [72] and Ni-CO [95] of D. vulgaris Miyazaki F. Note that in Ni-A a diatomic (oxygen based) ligand is in the bridging position X, the second oxygen blocking the free coordination site of the Ni opposite to the Cys549 ligand. In Ni-CO the extra CO ligand is attached axially to the nickel in the same position as the extra oxygen atom in Ni-A. Figure by courtesy of H. Ogata. Met. Ions Life Sci. 2, 279–322 (2007)


4.

295

SPECTROSCOPIC INVESTIGATIONS

In this section we review the most important and recent findings about the structure and function of the enzyme that were derived from spectroscopic methods. Since the enzyme has only little absorption in the visible region, optical techniques do not play a major role in hydrogenase research. The emphasis is therefore on EPR and FTIR techniques, but some results from other methods such as XAS will also be discussed. A separate section is devoted to modern electrochemical methods, these techniques have also been used in combination with FTIR, EPR, and other spectroscopies.

4.1.

FTIR Spectroscopy

By FTIR spectroscopy the active site of all redox states of the [NiFe] hydrogenase can be investigated. This technique allows for a sensitive detection of the vibrational frequencies of the three iron-coordinating diatomic ligands and was originally used to identify them as two CN and one CO by isotope labeling, where X-ray crystallography could not determine their identity with certainty [21,22]. CO and CN are σ-donor/ π-acceptor ligands that accept electron density from the iron and stabilize it in a low oxidation state (FeII). In addition, the strong ligand field favors the FeII low-spin (S 0) state. Changes of the redox state or of the ligand sphere of the hydrogenase active center affect the electron density at the iron and thereby the Fe-C bonds and the CO, C⬅N vibrational frequencies. Thus, FTIR spectroscopy can be used as a method to study the details of the electronic structure, including the EPR silent states, by monitoring the CO and CN frequencies [23–26,96–100]. A set of spectra for different redox states obtained for the enzyme from D. gigas is shown in Figure 6. The vibrational frequencies of the CO and CN ligands for three different catalytic [NiFe] hydrogenases (A. vinosum, D. gigas, and D. vulgaris Miyazaki F) are very similar (see Table 2). Small deviations can be attributed to slight differences in the amino acid surrounding of the active site that affect the vibrational frequencies, e.g., via hydrogen bonding or charge effects. FTIR studies were also performed on the hydrogenases from R. eutropha. The MBH showed spectra similar to those of the other catalytic [NiFe] hydrogenases, whereas for the SH a different pattern was found [101,102]. Here a third CN ligand was detected that is probably bound to the nickel and blocks the active site. The regulatory hydrogenase RH in the Ni-C state shows the same spectrum as the Ni-C state in the catalytic hydrogenases [103,104]. Recently, stopped-flow FTIR studies have been published by Albracht, Thorneley et al. [97,98] in which the reaction of the A. vinosum enzyme with H2, CO, and O2 was measured as a function of different temperatures and pH values. Met. Ions Life Sci. 2, 279–322 (2007)

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1946

0.002 O.D.

Ni-B 2090 2079

Ni-SIa 1934 1914

Absorbance

Ni-SIr 2075 2069 2055 2086

Ni-C

1952

2086

2073

Ni-R

1940 2073 2060

2150

2100

2050

2000

1950

1900

–1)

Wavelength (cm

Figure 6. FTIR spectra of various states of the [NiFe] hydrogenase of D. gigas obtained by electrochemical reduction of the Ni-B state. Reprinted with permission from [25]. Copyright 1997 American Chemical Society.

These investigations gave new insight into the activation process and the inhibition of the hydrogenase and the catalytic cycle. The combination of FTIR spectroscopy with an electrochemical cell (Figure 7) [25,26,42,99] allows monitoring of the electrochemically induced redox transitions between the intermediate states as function of pH, temperature and time. This has been beautifully demonstrated for different enzymes by several laboratories

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Table 2. IR absorption bands of the different redox states of [NiFe] hydrogenase from D. vulgaris Miyazaki F (DvMF) [26,176], A. vinosum (Av) [99], and D. gigas (Dg) [25]. Redox state

∼ νCO (cm1)

Av

Dg

1945 1943 1948 1910 1931 1950 1936 1921 1913

1947 1946 1950 1914 1934 1952 1940 1923 nd

DvMF Ni-A Ni-B Ni-SU Ni-SI r Ni-SIa Ni-C Ni-R Ni-R" Ni-R'

1956 1955 1946 1922 1943 1961 1948 1933 1919

∼ νCN (cm1)

DvMF 2084 2081 2075 2056 2075 2074 2061 2050 nd

∼ νCN (cm1)

Av

Dg

2082 2079 2088 2052 2073 2073 2059 2048 2043

2083 2090 2089 2055 2075 2073 2060 2050 nd

DvMF 2094 2090 2086 2070 2086 2085 2074 2065 nd

Av

Dg

2093 2090 2100 2067 2084 2085 2072 2064 2058

2093 2090 2099 2069 2086 2086 2073 2060 nd

nd not determined.

relative absorbance

1.0

pH 7.8 b

a 0.8

c Ni-B Ni-SI Ni-C Ni-R

0.6

0.4

h g

e

0.2

g

0.0 –600 –500 –400 –300 –200 –100

0

b f

1cm

a

100 200 300

potential (mV) vs. NHE

Figure 7. Spectroelectrochemical titration of the [NiFe] hydrogenase of D. vulgaris ∼ Miyazaki F at pH 7.8 and T 30C monitored by the νCO band in the FTIR spectrum (ready and active states only) [26]. The lines display Nernst fits with n 1. The midpoint potentials are directly obtained (at 0.5). On the right the electrochemical cell used for this work is shown [180], the arrow indicates the optical transmission path; (a) CaF 2 window mounted on (b) Plexiglas ring; (c) plastic body; (e) platinum counter electrode; (f) gold mesh working electrode; (g) rubber O-ring; (h) capillary connection to reference electrode.

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Table 3. Midpoint potentials of the different redox transitions of D. vulgaris Miyazaki F and of D. gigas [NiFe] hydrogenase measured at pH 8. Em (H) (mV) Species D. vulgaris D. gigasb D. gigasc

a

Ni-A/Ni-SU

Ni-B/Ni-SI

Ni-SI/Ni-C

Ni-C/Ni-R

Reference

96d 120

147 150 140

340 380 330

404 445 405

[176] [25] [177]

a

From FTIR data, the error lies in the range 5–10 mV. From FTIR data, the pH dependence of the 4 transitions (left to right) is 55, 36, 47 and 43 mV/pH unit. c From EPR data. d Value measured at pH 6.0. b

[25,26,42,99]. It is important to notice that the redox titrations followed by FTIR for the D. gigas enzyme are identical to those described earlier by EPR spectroscopy [12,25]. In this way the redox potential for all transitions of the enzymes from D. gigas [25], A. vinosum [99], and more recently D. vulgaris Miyazaki F [26] could be measured, the data are collected in Table 3. An example for such a titration is shown in Figure 7. It was shown that each transition is best fit by a Nernst equation with n 1 (single electron transition) and that the transitions are all pH-dependent, i.e. electron transfer is accompanied by transfer of a proton [25,26,42,99]. Recently, an extensive spectroelectrochemical study of the enzyme from A. vinosum has been published by Bleijlevens et al. [99] showing at least 13 redox intermediates. The conclusions derived from this work on the activation/deactivation process and the catalytic cycle are discussed in Section 6. However, some of the intermediates and transitions are not found in other enzymes and are therefore still under debate [33]. Spectroelectrochemical measurements have also been performed as a function of time (usually at reduced temperature) from which kinetic information of the various redox steps can be obtained (see, e.g., [25]). Such information is particularly valuable when combined with H/D exchange (isotope effect) and with studies of site directed mutants as recently presented by De Lacey et al. [100]. The spectroelectrochemical redox titrations on the [NiFe] hydrogenase (see Figure 7) have shown that it is difficult in many cases to generate a pure redox state; contamination with other states will frequently occur. This has important consequences for the results derived from other techniques on these enzymes. DFT calculations have been performed by Hall et al. in which calculated structural parameters (CO bond distances) were calibrated with measured IR stretching frequencies [105] (see also [106,107]). These calculations were used to

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propose the structures of the intermediates of the [NiFe] hydrogenase. Calculations of vibrational frequencies were also performed by Amara et al. [108], showing deviations from the experimental frequencies, in particular for νCO, that are typical for DFT methods (cf. also [109]).

4.2.

X-Ray Absorption Spectroscopy

X-ray absorption spectroscopy (XAS) is an important method for studying metalloenzymes since it is able to yield information about the oxidation state of the metal(s) (by XANES) and the number and type of ligands and their distances from the metal center (by EXAFS). Thus, XAS is able to provide electronic and structural data complementary to information obtained from X-ray crystallography and other spectroscopic methods. Several publications have addressed the structure of [NiFe] hydrogenase in recent years [27,110–116]. Ni-K-edge XAS spectra show that the reduction of the enzyme and the formal redox state of the Ni atom are not correlated. The data are consistent with the formal oxidation state of the Ni metal oscillating between NiIII in all EPR active states (Ni-A, Ni-B, Ni-C) and NiII in the EPR-silent states (Ni-SU; Ni-SI; Ni-R) [111,112]. Investigation of the photoconversion of the Ni-C to the Ni-L state shows that the photoproduct is slightly more reduced than Ni-C, but the observed edge shift is too small to justify description as a NiI state. Experiments on the reduced EPR-silent states using Ni L-edge XAS indicated that the NiII is in a high-spin (hs) state (S 1) [117]. A high-spin signal has not been observed yet by EPR spectroscopy, possibly because of a zero-field splitting of the nickel larger than the microwave quantum, which inhibits detection by conventional X-band (9 GHz) EPR spectroscopy. Here, high-field EPR (100 GHz) is required for a detection of the NiII high-spin state. In a recent publication by Bruschi et al. the Ni-SI and Ni-R states are treated by density functional theory [116]. The high-spin and low-spin states were found to be very close in energy so that slight structural alterations could already interchange the order of the two states. For Ni-SI a model was chosen in which the bridge between Ni and Fe is empty and for Ni-R a hydride is present. The authors noted that the structural features of the Ni-R model are compatible with X-ray data of the reduced hydrogenase. The EXAFS analysis of the D. gigas enzyme [111] in the oxidized state revealed two Ni–S bonds of about 2.2 Å, one or two Ni–S bonds of about 2.35 Å and a Ni–O bond of 1.91 Å length. These distances are close to those obtained from X-ray crystallography for similar enzymes (see Table 1 in [39]). For the reduced enzyme two Ni–S distances increased to 2.47 Å, but a Ni–O distance was still observed which disagrees with the X-ray structure. EXAFS data of the A. vinosum enzyme [112] suggest one O-ligand in Ni-A and Ni-B, two in Ni–SU, but loss of

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the oxygen in the more reduced states, for which also a shortening of the Ni–Fe distance from 2.85 to 2.5–2.6 Å was observed in agreement with crystallography. For the EPR-silent inhibited Ni-CO complex a nickel–carbon distance of 1.78 Å was found, showing that the exogenous CO binds to the nickel, probably blocking the open coordination site. This geometry is in excellent agreement with the recently published structure of the Ni–CO complex [95] (see Figure 5). For the regulatory hydrogenase of R. eutropha K-edge measurements have also been performed [115]. The EXAFS studies revealed a rather unexpected change in the coordination of the nickel. For the as isolated oxidized state a NiII(O,N)2S3 and in the H2 flushed state a NiIII(O,N)3X1S2 [X being either O, N or H] active site was proposed [115]. More experiments are required to solve the question of how the [NiFe] center of the RH can have a different coordination and ligand field and at the same time almost identical FTIR [104] and EPR spectroscopic properties [81] as the catalytic hydrogenases. For the soluble hydrogenase of R. eutropha XAS studies [118] revealed that the Ni is six-coordinated in the oxidized state (NiII(CN)O3S2) carrying only two Cys S and a CN ligand. Prolonged reduction resulted in a standardlike NiII(CysS) 4 site which could be further reduced to form the Ni-C state (NiIIIH). The specific coordination of the oxidized form obviously stabilizes the EPR-silent NiII state. The Ni-C state has been proposed to be inactive in hydrogen cleavage.

4.3. EPR Spectroscopy EPR spectroscopy is uniquely suited to obtain information about the EPR active redox states of the [NiFe] center. These include the oxidized ‘ready’ (Ni-B) and ‘unready’ (Ni-A) states, the Ni-C state, as well as the light-induced Ni-L states, and the EPR-active form of the CO inhibited state. EPR was used to identify these states and characterize them by their specific EPR signals obtained from frozen solutions of the enzyme. Each state shows a rhombic g tensor with three characteristic components gx, gy, gz. These signatures were also used as markers for redox titrations, carried out as function of pH, temperature and time [18,119,120]. Furthermore, several oxidation states of the iron–sulfur clusters (both [3Fe4S] and [4Fe4S]) could be monitored by this technique [14]. In the oxidized state the [4Fe4S] 2 clusters are EPR-silent but the [3Fe4S] can be observed at low temperature. The signal is superimposed to that of the Ni-A/B state. This cluster has been characterized by high-field EPR yielding the complete g tensor (g1 2.026, g2 2.017, g3 2.011) [121]. In the reduced state (Ni-C) the [4Fe4S] 2 clusters eventually also become reduced. At low temperature (10 K) the [NiFe] center magnetically interacts with the proximal [4Fe4S] cluster which leads to a splitting of the g tensor components of the Ni-C [122]. From the analysis of these spectra a set of structural and magnetic (dipolar and exchange) parameters was deduced [123]. For the light-induced Ni-L signal(s) a splitting was also observed Met. Ions Life Sci. 2, 279–322 (2007)


301

at low temperature and led to structural information [124]. Elsässer et al. used pulsed electron–electron double resonance (PELDOR) to study the interaction between the [NiFe] center (Ni-B/A) and the [3Fe4S] center. This analysis also gave insight in the spin coupling of the [FeS] cluster [125]. Data obtained from EPR also include information on the spin and valence states of the nickel and its coordination environment in the various redox states. A powerful approach is combination of EPR with isotope labeling of the samples that led to identification not only of the nickel (61Ni) in the enzyme [10], but also of oxygen (17O) and nitrogen (15N) ligation in [NiFe] hydrogenase and, selenium (77Se) in case of the [NiFeSe] hydrogenase [126–128]. The early EPR work before the first X-ray crystallographic structure has been reviewed in detail by Albracht in 1994 [13]. During the last decade additional EPR experiments have been performed, in particular on single crystals of the [NiFe] hydrogenase from Desulfovibrio vulgaris Miyazaki F, which gave very precise information about the magnetic tensors. The approach, illustrated in Figure 8, yielded g-tensor components and orientations for Ni-A and Ni-B [129] and also for Ni-C and Ni-L [130]. The EPR-active Ni-CO state has been investigated so far only in frozen solution [131]. These data were compared with those derived from density functional (DFT) calculations obtained for geometry optimized models of the [NiFe] site (see Table 4). In these calculations some of the ligands were varied, in particular the bridging ligand X. The final analysis of all data yielded the following conclusions about the active site [17,132]:

•

Ni-A, Ni-B, and Ni-C are formally all NiIII species with a spin state S 1/2 and a dz ground state. The dz2 orbital is oriented along the molecular z-axis (gz-axis, gz ⯝ ge) pointing to the open coordination site of the Ni (see Fig. 4B). 2

Table 4. Comparison of experimental and calculated g-tensor values for various states of [NiFe] hydrogenase (for details see [133]). Calculationb

Experiment a State Ni-COc Ni-L Ni-C Ni-B Ni-Ad

gx

gy

gz

gx

gy

gz

2.12 2.30 2.20 2.33 2.32

2.07 2.12 2.14 2.16 2.24

2.02 2.05 2.01 2.01 2.01

2.11 2.26 2.20 2.21 (2.36)

2.06 2.10 2.10 2.17 (1.95)

2.00 2.05 2.00 1.98 (1.84)

a

Experimental data for Ni-CO from [18], all other data from [132]. Calculated data from [133]. c Ni-CO: CO bound end-on to nickel. d For Ni-A a µ-oxo bridge was used in the calculation [133]. The result suffered from orbital degeneration leading to large effects on the calculated g-values [140]. b

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2.4

T=80 K frozen solution

g-value 2.2 2.1 2.0

1.9 Ni-A Ni-B

(B)

w θ

θ

v

u

0° c

20°

θ

(A)

w

b

y

x z

a

40° 60°

B0

v

u

EPR spectra were recorded after each rotation of the crystal about ∆ θ.

80° Angular Dependence of the g2 - Values

100° 5.2

120°

(C)

Ni-B

5.0 g2

140° 160°

4.8 4.6

IV

180°

II I

4.4

III

4.2 288 304 320 336 352 368 B0[mT]

0

30

60 90 120 150 Rotation Angle θ [deg]

180

Figure 8. (A) Crystal rotation in the laboratory axis system (u, v, w), also shown are the crystallographic (a, b, c) and the molecular (g tensor) axes (x, y, z). (B) EPR spectra (X-band) obtained for a mixture of the oxidized states (NiA/Ni-B 1:2) in the [NiFe] hydrogenase of D. vulgaris Miyazaki F for a frozen solution and single crystal (rotated in steps of 5). (C) Angular dependence of the g2 value (g Zeeman splitting factor) for the dominant Ni-B state (solid lines) and the Ni-A (grey dots). Four sets of lines are obtained for each state which are related to the four sites (I to IV) in the unit cell (P212121 space group of the crystal). The evaluation yields the g tensor components and angles with respect to the crystallographic axes. For further details see [129].

• • • • •

Ni-L is best described by a dz2 ground state with a small admixture of d x2 –y2. Nickel is redox active, it oscillates between NiIII and NiII. Iron is always in the FeII low-spin (diamagnetic) state, probably caused by the strong ligand field (CN, CO ligand). Iron is not redox active. The bridging ligand X between Ni and Fe (Fig. 4B) is changed in the reaction cycle; best agreement between experiment and calculation is found for the following assignments Ni-B: µ-OH, Ni-C: µ-H, Ni-L: empty; for Ni-A a final conclusion could not be reached (see Section 6 and [143]). The Ni–CO state is derived from Ni-L with a formal NiI ground state and CO axially bound to the Ni [133].

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303

Ni-C CN

Ni H-

Fe CO CN

Figure 9. Contour plot of the unpaired spin density distribution (0.005 e/a03), DFT (BLYP/DZVP) of a model mimicking the Ni-C (NiIIIFeII) state. Note that the cysteine residues are modeled by CH3CH2S residues. The bridge is occupied by X H. The total Mulliken atomic spin densities are at Ni: 0.51, Fe: 0.01, X(H): 0, bridging S: 0.29, terminal S: 0.10 [109].

The calculated spin density distribution of the models for the active site show that nickel carries ⬃50% of the spin, the rest is delocalized over the sulfur ligands (see Figure 9). Only a vanishing amount is at the iron, which is in agreement with the small 57Fe hyperfine coupling constant [134]. The calculations exclude an SO ligand at the Fe, which had been proposed earlier [88], and support ligation by CN and CO. Replacement of the oxygenic species in the Ni–Fe bridge by a sulfur species also led to poorer results [132]. The results of the g tensor analysis were recently further corroborated by detection of the 61Ni hyperfine tensors measured on 61Ni labeled [NiFe] hydrogenase from D. vulgaris Miyazaki F [130]. Further corroboration of the proposed structural models of the [NiFe] hydrogenase active site came from the application of advanced EPR techniques that are able to resolve the electron nuclear hyperfine (hf) and nuclear quadrupole (nq) interactions of the magnetic nuclei of the ligands. Such experiments are discussed below.

4.3.1.

The Intermediate Ni-C State

The Ni-C state was investigated by ENDOR spectroscopy in the early 1990s, and it was found that a very strongly coupled exchangeable proton was present with a hyperfine coupling constant of about 20 MHz [135,136]. This proton disappeared upon illumination of the sample at low temperature (100 K) and formation of the light-induced Ni-L states, but upon thermal annealing, the signal reappeared Met. Ions Life Sci. 2, 279–322 (2007)

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[135]. A determination of the complete hyperfine tensor of this proton has been performed only recently by ENDOR and HYSCORE spectroscopy, first on the regulatory hydrogenase of R. eutropha [137] and then for the catalytic one from D. vulgaris Miyazaki F [138]. Upon solvent exchange to D2O and reduction with D2 gas, a very rich structure appeared in the deuterium region of the HYSCORE spectrum (see Figure 10). Analysis of these signals [137] revealed that they result from one exchangeable proton/deuterium, the same as observed earlier [135,136]. The HYSCORE spectra measured after illumination of the Ni-C sample showed that the deuterium signals indeed disappeared (Figure 10). The complete hyperfine tensor was elucidated from these experiments and compared with that from 1 H ENDOR and with those of protons in the active site as found in DFT calculations. The comparison uniquely positioned the proton (or the respective deuteron) as a bridging ligand between nickel and iron and it was assigned as a formal

2H 4

HYSCORE

gz

3 MHz

−

R RS

ν2

2

Ni H

R S S

CO Fe

2 –

H

RS

CN CN

1

Ni-C 0

1

2 MHz

3

4 20 min white light

T < 100 K 2H

4

gz

3 MHz

HYSCORE 2–

R

ν2

RS

H

2

Ni

R S S

RS

1

Ni-L 0

1

2 MHz

3

4

CO Fe

CN CN

2 + H (base)

Figure 10. HYSCORE (four-pulse ESEEM) spectra [137] of the Ni-C and Ni-L (Ni-L2) states in the deuterium region obtained after H/D exchange in the regulatory [NiFe] hydrogenase from R. eutropha (RH). Note the disappearance of the 2H hyperfine coupling upon illumination of Ni-C. A structural model is shown on the right (removal of hydride bridge as H). Met. Ions Life Sci. 2, 279–322 (2007)


305

hydride (H) [137,138]. The presence of a hydride bridge was already postulated earlier by DFT calculations [108,133,139–141]. Upon illumination the bridging hydrogen leaves as a proton, H. The most likely acceptor for the proton, given the reversibility upon annealing of the sample and the observation that the Ni-L states have the same spectroscopic characteristics in many hydrogenases, is a conserved nearby amino acid.

4.3.2.

The Oxidized States

The identity of the bridging ligand in the oxidized states could not be fully established by EPR on the native enzyme. In an early experiment 17O labeled oxygen [90] was used to show that this ligand is oxygen-based in A. vinosum hydrogenase in both redox states. However, the observed line-broadening after treatment of the sample with molecular 17O2, did not show whether the molecule binds as molecular O2, or whether it has already been partially reduced, as recently suggested by electrochemical data [93,94]. An argument that O2 cannot be bound as such near the active site, is due to the triplet ground state of O2 which would lead to significant spin–spin interaction. 17O ENDOR experiments were performed on Ni-A of D. gigas hydrogenase treated with H217O following a reduction/oxidation cycle [91]. These experiments confirmed that the bridging ligand can be derived from solvent water and is indeed oxygen-based. An identical experiment for the ready state Ni-B has not yet been performed. Single-crystal 1H ENDOR experiments have recently been carried out on D. vulgaris Miyazaki F hydrogenase in the oxidized state. The signals of one exchangeable proton were observed for the Ni-B state that could unequivocally be assigned to the bridging ligand [142]. The ENDOR experiments also allowed determination of the position of the hydroxyl (OH) ligand in the ready oxidized (Ni-B) state. Due to the great similarity of the catalytic hydrogenases this conclusion should also hold for the other [NiFe] hydrogenases. However, it cannot be excluded that under certain conditions a sulfur-derived ligand (e.g., SH, see Section 3) can occupy the bridging position, which should show different spectra. For the Ni-A state final conclusions on the type of bridging ligand could not yet be drawn. Recent HYSCORE experiments and H/D exchange experiments show also for Ni-A the presence of an exchangeable proton [143]. A large variety of DFT calculations have been performed earlier on the oxidized states of the enzyme showing differing results. Pavlov et al. [144] postulated high mobility of CN ligands moving from a terminal to a bridging position. An O2– bridge has been suggested in the unready state by Amara et al. [108]. The ready state had an empty bridge in the work of Niu et al. [105]. DFT calculations by Stein et al. suggested an OH bridging ligand for the ready state and an O2– for the unready state [139,140]. Stadler et al. found that an OH bridge is most likely for both the unready and ready states [145]. Recent DFT calculations [142,143] Met. Ions Life Sci. 2, 279–322 (2007)

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indicate as possible candidates an OH (in a conformation different from that in Ni-B) or an OOH ligand. The latter hypothesis is compatible with recent X-ray crystallographic data [71] [72].

4.3.3.

Interaction with the Protein Environment

The active [NiFe] center interacts with the protein via Coulomb or hydrophobic interactions or hydrogen bonds. A H-bond has been postulated from a highly conserved histidine residue (His-88 in D. vulgaris Miyazaki F) that is formed to the axial cysteine (Cys-549 in D. vulgaris Miyazaki F) at the nickel which carries significant spin density (Figure 9). Three-pulse and four-pulse ESEEM (HYSCORE) experiments yield a significant 14N hyperfine coupling and a 14N nuclear quadrupole coupling characteristic for Nε-H of histidine. 15N labeling of His [146] finally assigned this residue in D. vulgaris Miyazaki F. It is assumed that the histidine plays a role in fine tuning the functional properties of the active site of the [NiFe] hydrogenase. Interestingly, this His is absent in the regulatory hydrogenase of R. eutropha, for which a similar interaction could also be detected by ESEEM after incorporation of a His residue in place of glutamine near the active [NiFe] site [81]. The results obtained experimentally were supported by DFT calculations of the respective nuclear quadrupole coupling constants [146].

5.

ELECTROCHEMISTRY

Electrochemical titrations using a spectroelectrochemical cell have been performed to study the redox transitions of the [NiFe] hydrogenase and their pH and temperature dependence [25]. The characteristic IR bands of the CO/CN at the iron have been monitored. The field has recently been reviewed by Best [42]. The data were compared and assigned to specific states in the redox cycle of the enzyme which were created by gas treatment with appropriate electron acceptors/donors [97,98]. These experiments played a central role for identification and characterization of the various intermediates and were instrumental in the development of a reaction mechanism (see Section 6). Here another electrochemical technique complementary to spectroscopy will be discussed, i.e. protein film voltammetry, which was recently applied successfully to hydrogenase research [31–33,92–94,147–153] (see Figure 11). In protein film voltammetry a small amount of enzyme is applied to a pyrolytic graphite edge (PGE) working electrode under unaerobic conditions to study the reaction with added gases (H2, N2, O2, CO) or gas-saturated solutions. For effective mass transport to the surface the electrode is rotated and the potential controlled and measured via two other (counter and reference) electrodes. For further details of the technique the reader is referred to [147]. Met. Ions Life Sci. 2, 279–322 (2007)


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Figure 11. Schematic view of a protein film voltammetry experiment on a hydrogenase attached to a rotating PGE electrode by courtesy of F. Armstrong, Oxford University, UK.

It has been shown that with hydrogenase a highly active robust film can be obtained, allowing the enzyme to be cycled over extended time periods. For the A. vinosum enzyme at 30C turnover rates of up to 10000 H2 molecules per second are reached [92,147], which is much higher than those obtained in other activity assays. Proton reduction has been found to be slower. With this enzyme electron flow through the iron–sulfur clusters, various aspects of the catalytic cycle (electron–proton coupling, catalytic bias), oxidative inactivation with/without O2, and reductive activation were measured. Thereby a mechanism for the activation was established and the differences between the inactive states (Ni-A, Ni-B) could be described. Furthermore, [NiFe] hydrogenases from different organisms were studied. Of particular interest are those from R. eutropha [154] which are not inhibited by CO and show a small oxygen sensitivity in contrast to most other catalytic [NiFe] hydrogenases. In the following the major results are summarized (see also [32,92]). (i) The mechanism of transformation between states of the catalytic cycle has been elucidated by kinetic studies. Here inactivation and activation were investigated by stepping in and out of the potential region for H2 oxidation activity and following the time course of the current as function of potential which is directly related to the catalytic turnover rate. It was found that the rate of oxidative inactivation is independent of potential, but dependent on pH, i.e., the first stage must be a chemical rather than an electron-transfer event. Obviously, to inactivate the hydrogenase the bridging ligand X (a hydroxide) has to enter the site before oxidation of nickel NiII → NiIII takes place. The activation step is, however, strongly dependent on potential, showing that in this case the reduction of nickel precedes the subsequent reaction (Scheme 1) [152]. In this activation/inactivation mechanism the proton entering the enzyme in the Ni-SI state [25,26,155] is Met. Ions Life Sci. 2, 279–322 (2007)

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NiIIIOH–FeII Ni-B

+ e– –

e–

+ H+

NiIIOH–FeII Ni-SIr

– H+

H2O FeII NiII Ni-SIa

(1)

Scheme 1.

related to a protonation of the OH ligand which leaves the active site as water molecule leading to the catalytically active Ni-SIa state. (ii) Reaction of active [NiFe] hydrogenase with O2 leads to two stable products, the unready Ni-A and the ready Ni-B state. Furthermore, it was found that other nonrecoverable products are formed. The ratio depends strongly on the reducing conditions of the solution when O2 is injected. Under reducing conditions (0 mV, H2 present) injection of O2 only produces the Ni-B state in the A. vinosum enzyme. Under oxidizing conditions (200 mV, under N2) more than 80% of the enzyme gets in the Ni-A state [93,94]. Based on these results and the available EPR and structural data Armstrong and coworkers proposed that when electrons are abundant, reduction of O2 proceeds rapidly by four electrons to two water molecules (one effectively retained as OH). Smoothly the Ni-B state is formed and no further intermediates are trapped (see Scheme 2, reaction (2)). However, when electrons are not abundant O2 reacts only to the level of peroxide, reaction (3), thereby creating a powerful and potentially damaging oxidant to attack the active site producing Ni-A and other products with oxidized cysteines (SO groups), reaction (4). Based on the recent X-ray structures of the Ni-A state [39,72] this intermediate indeed seems to carry a peroxide or hydroperoxide ligand in the bridge between Ni and Fe (see Fig. 5). In both X-ray structures the oxidation of cysteine sulfurs is also evident. These results are of great importance for understanding the inhibition of the enzyme by molecular oxygen which limits the use of hydrogenases for biotechnology combining light-induced water oxidation (producing oxygen) and hydrogen production by hydrogenases.

–

Ni II Fe II + O2 + 3e– + 3H + Ni-SIa

Ni III OH Fe II + H2O Ni-B

(2)

Ni II Fe II + O2 + e– + H + Ni-SIa

Ni III OOH – Fe II Ni-A

(3)

Ni II Fe II + O2 + e– + 2H + Ni-SIa

Ni III Fe II S=O Ni-A


or

Ni III Fe II + H2O [O] Ni-A

(4)


V

(A)

309

(B) 5 H2 Hydrogenasecoated anode

Laccasecoated cathode H+ O2 H2O H2

4

Power / µW

Air

Load

3 2 1 0 0.1

10

1000

100000

Load / kΩ (logarithmic scale)

Figure 12. (A) Hydrogen/oxygen fuel cell without a membrane using PGE electrodes with enzymes as electrocatalysts. The anode is coated with the MBH of R. eutropha, the cathode with laccase from white rot fungus. (B) Power output as a function of applied load curve for the set-up shown in (A). The squares are for the oxygen-insensitive MBH, the circles for an oxygen sensitive hydrogenase (from A. vinosum). Reprinted with permission from [154]. Copyright 2005, Natl. Acad. Sci. USA.

(iii) It was found that different enzymes at the PGE electrode are highly active under anaerobic conditions at elevated temperatures and are able to catalyze hydrogen cycling (H2 L 2 H 2 e) in either direction [147,148]. They are also functioning at oxidizing potentials (⬵400 mV). The high electrocatalytic activity might allow the use of hydrogenase on PGE as alternative to expensive Pt electrodes in fuel cells [6,92,154], in particular when enzymes are used that are CO and oxygen insensitive. This was recently demonstrated by Armstrong and coworkers [154] who constructed a working electrocatalytic cell (laccase coated cathode, hydrogenase coated anode) (see Figure 12). For this purpose the membrane-bound hydrogenase from R. eutropha was used that is rather insensitive towards oxygen.

6.

HYDROGENASE FUNCTION AND THE CATALYTIC CYCLE

Based on the various data described in the preceding sections, a large number of (possible) intermediates of the reaction cycle of [NiFe] hydrogenase could be identified and structurally characterized. In Figure 13 the relationship between the different states is shown that is based on electrochemical studies of different catalytic hydrogenases [40,42,99]. The enzyme cycles between NiIII and NiII states as indicated, the iron (FeII) is not changing the oxidation state. The FeII is always low spin (S 0, diamagnetic) Met. Ions Life Sci. 2, 279–322 (2007)

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FeII[O]NiIII

FeII(OH–)NiIII

active

inactive Ni-A*

Ni-B* e–, H+

e–, H+(?) +

FeII[O]NiII

Ni-SU

FeII(OH–)NiII

Ni-SIr

+/–H

(H2O)

–/+H2O

FeII NiII

Ni-SIa

FeII NiII

e–, H+ FeII(H–)NiIII

Ni-C*

e–, H+

FeII(H–)NiII

Ni-R

Figure 13. Schematic overview of the different states of the catalytic [NiFe] hydrogenase based on spectroelectrochemical experiments monitoring the IR bands of CO/ CN at the Fe. The EPR detectable states (S 1/2) are denoted by an asterisk. For the various intermediates the oxidation number is given for Ni and Fe and also the type of bridging ligand between them. Further details are described in the text, see also [42]. Different nomenclatures are used in the literature for the different states, e.g. Ni-A Ni*u ; Ni-SU Niu-S, Ni-B Nir*; Ni-SI r Ni-SI I or Ni r-S; Ni-SIa Ni-SI II or Nia-S, Ni-C Nia-C*, Ni-R Ni-SR. Sometimes the states are additionally denoted by the ∼ vibrational frequency of νCO [99].

and NiIII has a d1z2 ground state (S 1/2) [132]. The EPR-silent states (NiII) have been proposed to exist in high-spin (S 1) states by XAS spectroscopy [117]. By DFT calculations, the singlet and triplet states of the NiII were found to be very close in energy [156] and spin cross-over was suggested to be possible at room temperature. In the unready oxidized Ni-A state the bridging ligand [O] may be an OH or an OOH [39,71,72], it is likely that the ligand is retained in Ni-SU, but little structural information is available about this state. The activation/inactivation process involving states Ni-A, Ni-B and Ni-SI has been discussed above (Schemes 1, 2). Further details are found in [40]. Recently a new EPR-silent inactive state (Ni-S) was described assigned to an S-bound species [99]. The inactive ready oxidized state Ni-B has been conclusively shown to contain a OH bridge, the metal redox states could also be clarified [132,142]. A reductive step leads to the Met. Ions Life Sci. 2, 279–322 (2007)


311

Ni-SI state which is present in an acid-base equilibrium Ni-SIr L Ni-SIa. It has been proposed [32,99] that the bridging ligand OH is protonated to form H2O in this process. In A. vinosum hydrogenase an inactive state could be trapped at low temperature (2C) [157]. It has been speculated that here the water is still present near the active site and obstructs the binding of H2. This state could not be observed in other hydrogenases so far [25,26]. At higher temperatures the water is removed and a highly active state, Ni-SIa, is formed that readily reacts with H2, O2, and CO and is thought to be part of the catalytic cycle. With H2 this state forms Ni-C which has been shown to carry a hydride bridge between NiIII and FeII, the state is EPR active and considered to be a central intermediate in the catalytic cycle [132,158,159]. The observed two spectroscopic forms of Ni-C [99] have been attributed to different redox states of the proximal [4Fe-4S] cluster. Further reduction of Ni-C leads to the fully reduced Ni-R state which still carries the hydride [39,42,99]. This state exists in different subforms. It has been proposed that these forms represent different protonation states of Ni-R [99,100] but could also be due to different spin states of the NiII and/or conformations of the active site [26,39]. In this model (Fig. 13) the catalytic cycle involves the Ni-SIa, Ni-C and Ni-R states [25,97]. Redox states of the [NiFe] hydrogenase not shown are the Ni-L states (Ni-L1 and Ni-L2) that are formed upon illumination of Ni-C at different temperatures. These states have been described initially as formal NiI states with a mixed dz2 / d x2y2 ground state (EPR active, S 1/2). In Ni-L the bridging hydride is (reversibly) transferred to a nearby base as a proton [130,137]. The existence of NiI states in the enzyme has, however, been questioned [112]. It has been speculated [39,160] that Ni-L-like states could occur as transients in the catalytic cycle. Furthermore, CO-inhibited states exist that can be formed by reacting either Ni-SIa or Ni-C with CO, which are EPR-silent and EPR-active, respectively. The latter has been described as a NiI-CO species that, upon illumination yields the Ni-L state. A more detailed description of the various states and their interconversion can be found in [39,40,42,99,161]. Here the states of the active [NiFe] site have been discussed in some detail. A discussion of states of the [FeS] clusters during activation and the catalytic cycle is found in [13]. With all data available from the methods described in this chapter, first attempts have been made to establish a working model for the actual enzymatic reaction cycle of the [NiFe] hydrogenase. The first group to propose a model was Siegbahn et al. [144,162]. Others who have proposed mechanisms are Hall et al. [105], De Gioia et al. [156,163], Stein and Lubitz [133], and de Lacey et al. [40]. An overview of the similarities and differences of these mechanisms is given in a recent review [43]. The states involved in the catalytic cycle are believed to be Ni-SIa, Ni-C and Ni-R, which are interconverted by one electron/one proton equilibria. This has been shown by George et al. [97] and Kurkin et al. [98] with redox titrations and Met. Ions Life Sci. 2, 279–322 (2007)

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kinetic assays. The states are also active in H/D exchange [25]. The Ni is probably the site of H2 binding since the gas channel ends close to the Ni open coordination site, the binding of the inhibitor CO is at Ni and the putative OOH ligand in the inactive Ni-A state is also blocking this site. In the next step base-assisted heterolytic cleavage of the H2 molecule should take place, leading to a bridging hydride species. One of the candidates for acting as a base is the terminal cysteine (Cys530 in D. gigas [43]) that has a high temperature factor in crystallographic studies, indicating conformational flexibility and a possible role in proton transfer. Alternatively, a water molecule bound to the Fe has been proposed to act as base [133]. Ni-C is a NiIII species, thus formation of the hydride causes release of an electron to the proximal [4Fe4S] cluster, which could be coupled with the release of the first proton. The final step would then be the release of another proton and electron to give rise to the initial Ni-SIa state with an open bridge, ready for the next turnover. This last step could be preceded by a relocation of the proton from the bridge to the terminal cysteine, creating a state that resembles Ni-L [40]. Two proposed mechanisms are shown in Figure 14 which are based on spectroscopic results and DFT calculations [43,133,156,163]. In both cycles H2 is initially attached to Ni. In mechanism A, cysteine acts as base and the hydride is formed in the Ni-R state before electron release leads to Ni-C which is then converted back to the initial Ni-SIa state. In B [133] the H2 is polarized at a NiIII and the proton is accepted by water bound to Fe. Release of H3O leads to the hydride carrying Ni-C state which is converted by a second proton and electron to Ni-R. Here the proton is bound either by Ni itself or by the terminal cysteine [133]. Release of H2 and introduction of H2O completes the cycle. Unfortunately, even with the wealth of spectroscopic data available, especially for the EPR active states, a final conclusion on the exact mechanism has not yet been possible.

7.

CONCLUSIONS AND OUTLOOK

Since the review about [NiFe] hydrogenase appeared in the Metal Ions in Biological Systems series in 1988 [164] much additional information has been obtained about this enzyme from genetics, structural biology, spectroscopy, electrochemistry, and synthetic and theoretical chemistry. This progress has been described in the present review. Important aspects of the enzyme activation, inactivation/inhibition, and the catalytic cycle have been obtained and we are close to an understanding of the reaction mechanism of this important enzyme, although specific aspects are still poorly understood and controversial. Much is still to be learned in the field of molecular biology and genetics. Concerning the mechanism it is still not entirely clear which of the many ‘spectroscopic states’ are really involved in the enzyme’s activation/deactivation and cycling. Is Ni or Fe the place of H2 activation, which redox state of Ni aids in the heterolytic H2 cleavage, and is the base required in this process simply a cysteine ligand or is it Met. Ions Life Sci. 2, 279–322 (2007)


313

Ni-R 2− R

(A)

R

RS

H2

CO FeII CN

H

RS

2−

S S

NiII

CN

H

R R

RS NiII

S S

–e–

CO FeII CN

RS

−

CN

R

Ni-SIa

R

RS –e–,

–2H

+

S S

NiII

CO FeII CN

H

RS

CN

H

Ni-C

(B)

−

R R

RS

NiIII RS H2, –e–

S S

H2O

CN

-H3O+ 2−

H

R R

FeII

H

2−

NiII

CO

S

CN

Ni-SIa

RS

S

R

CO

R

RS

FeII

NiIII CN

RS

H2O

H

RS

CN

S S

CO FeII CN CN

Ni-C 2−

+

–2e–, –2H , H2O

R R

RS NiII RS

S S H

H

+

H , e–

CO FeII CN CN

Ni-R Figure 14. Two models showing the catalytic cycle of [NiFe] hydrogenase based on DFT calculations (A, B). For details see text. Adapted from [43] by permission of Elsevier.

indeed a water molecule, or another base? Furthermore the possible role of highspin NiII in the mechanism must be investigated. Little is known of the details of the protein’s function, e.g., in forming the binding pocket for the [NiFe] complex, the hydrophobic hydrogen channel and the electron and proton transport chain. Met. Ions Life Sci. 2, 279–322 (2007)

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Furthermore, no synthetic models exist to date that mimic the reaction performed by the [NiFe] hydrogenase. The [NiFe] and [FeFe] hydrogenase are highly active in the production and consumption of hydrogen and are therefore central research targets for a future biologically based hydrogen technology. Early experiments with the aim of combining hydrogenase as a producer of H2 with photosynthetic proteins that split water and produce O2 have already been performed in the 1970s [165] (see also [35]) and are still going on in many laboratories. Pertinent problems are the limited stability of the biological components, the low energy yield of the chloroplasts and the oxygen sensitivity of most hydrogenases. However, the process – that is actually performed in nature by certain classes of green algae and cyanobacteria [166] – stimulated researchers to study the mechanisms and details of the molecular and electronic structure of all the components involved, including the hydrogenases. If the essentials of the enzymes involved in the catalysis of hydrogen conversion are better understood, it may be possible to solve the stability and activity problems and use the organisms or the isolated enzymes in biotechnology [167–170] or build functional, more stable synthetic analogs, which could be used in devices for clean energy production [38,171–175].

ACKNOWLEDGMENTS We would like to acknowledge the many important contributions of all doctoral students, postdocs, and external collaborators, whose names appear in the respective references. We are especially grateful to B. Plaschkies and B. Deckers for typesetting the manuscript and preparing the figures. The work was supported by the Deutsche Forschungsgemeinschaft and the Max Planck Society.

ABBREVIATIONS DFT ENDOR EPR ESEEM EXAFS FTIR hyp HYSCORE MBH PELDOR PGE RH

density functional theory electron nuclear double resonance electron paramagnetic resonance electron spin echo envelope modulation extended X-ray absorption fine structure Fourier transform infrared hydrogenase pleiotropic hyperfine sublevel correlation (spectroscopy) membrane bound hydrogenase pulsed electron-electron double resonance pyrolytic graphite edge regulatory hydrogenase

Met. Ions Life Sci. 2, 279–322 (2007)


SH TAT XANES XAS

315

soluble hydrogenase twin arginine transport X-ray absorption near-edge spectroscopy X-ray absorption spectroscopy

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8 Methyl-Coenzyme M Reductase and Its Nickel Corphin Coenzyme F430 in Methanogenic Archaea Bernhard Jaun1 and Rudolf K. Thauer*2 1

Organic Chemistry ETHZ, ETH Hönggerberg HCI E317, CH-8093 Zürich, Switzerland <[email protected]> 2 Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Strasse, 35043 Marburg, Germany

1. INTRODUCTION 2. STRUCTURE AND PROPERTIES OF COENZYME F430 2.1. Structure of F430, of F430 Isolation Artifacts, and of a Natural F430 Derivative 2.2. Properties of Ni(II)F430 and Nonplanarity 2.3. Reduction of Ni(II)F430 to Ni(I)F430 and the Properties of Ni(I)F430 2.4. Oxidation of Ni(II)F430M to Ni(III)F430M and the Properties of Ni(III)F430M 2.5. Methyl-Ni(II)F430 and Methyl-Ni(III)F430 2.6. Model Reaction for Methyl-Coenzyme M Reduction with Coenzyme B to Methane 2.7. Models for Coenzyme M in the Active site of Methyl-Coenzyme M Reductase 3. MOLECULAR PROPERTIES OF METHYL-COENZYME M REDUCTASE 3.1. Crystal Structures of MCRox1-silent and MCRsilent Metal Ions in Life Sciences, Volume 2 © 2007 John Wiley & Sons, Ltd


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3.2. MCRred1 with Penta-Coordinated Ni(I)F430 3.3. MCRred1/2 with Coenzyme M Coordinated with the Sulfur of Its Thiol Group to Ni(I) of F430 in One of the Two Active Sites 3.4. MCRBPS with a Ni(III)-Alkyl Bond as One of Three Resonance Structures 3.5. MCRox1 with Coenzyme M Coordinated via the Sulfur of Its Thiol Group to Ni(III) of F430 as One of Two Resonance Structures 3.6. MCRox1-silent, MCRred1-silent and MCRsilent with Hexacoordinated Ni(II)F430 3.7. Methyl-Coenzyme M Reductase from Methanotrophic Archaea CATALYTIC PROPERTIES OF METHYL-COENZYME M REDUCTASE 4.1. The Reversibility of the MCR-Catalyzed Reaction 4.2. The Catalytic Efficiency of Methyl-Coenzyme M Reductases 4.3. The Catalytic Mechanism of Methane Formation 4.4. Indications for a Dual-Stroke Engine Mechanism 4.5. The Anaerobic Oxidation of Methane ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

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INTRODUCTION

In 1979 it was discovered that Methanothermobacter marburgensis (then named Methanobacterium thermoautotrophicum strain Marburg) requires nickel for growth on H2 and CO2 as sole energy and carbon sources (4H2 CO2 L CH4 2H2O; CO2 2H2 L [CH2O] H2O) [1]. This was quite unexpected since at that time urease was the only enzyme known to contain nickel and since the methanogenic archaeon did not require urea for growth [2]. M. marburgensis was subsequently found to contain the following eight nickel enzymes, the evidence for this being based both on biochemical and genome sequence information [3–6]: Methyl-coenzyme M reductase isoenzyme I (McrABG) and isoenzyme II (MrtABG). The two cytoplasmic isoenzymes contain the nickel corphin coenzyme F430 as prosthetic group (for structure of F430 see Figure 1) [4]. McrABG and MrtABG are synthesized under different growth conditions [7–10] and have different kinetic properties [11]. They are both involved in the reduction of CO2 with Met. Ions Life Sci. 2, 323–356 (2007)

METHYL-CoM REDUCTASE AND Ni CORPHIN COENZYME F430

325

COOH

905 Da O CH3

H HN H3C

3

H2NOC

1 20

H

19

5

N

N

COOH 10

N i+ N

N

12

18

15

13

COOH

HOOC 173 172

O COOH

Figure 1. Structure of coenzyme F430, the prosthetic group of methyl-coenzyme M reductase from methanogenic archaea. View from the β -face.

H2 to methane at the oxidation level of methanol (methyl-coenzyme M) by catalyzing the reduction of methyl-coenzyme M (CH3-S-CoM) with coenzyme B (HSCoB) to methane and the heterodisulfide CoM-S-S-CoB (Figure 2). The presence of nickel in F430 was discovered by Diekert et al. [12] and Whitman and Wolfe [13] and the presence of F430 in methyl-coenzyme M reductase by Ellefson et al. [14]. F420 -reducing [Ni-Fe]-hydrogenase (FrhABG). The cytoplasmic enzyme is composed of three different subunits of which FrhA harbors the binuclear [Ni-Fe] center [15], FrhG is an iron–sulfur protein and FrhB an iron–sulfur flavoprotein. The hydrogenase is involved in CO2 reduction with H2 to methane at the oxidation levels of formate (methenyltetrahydromethanopterin) and formaldehyde (methylenetetrahydromethanopterin). Frh is not synthesized under nickel-limiting growth conditions, under which its function is taken over by the iron–sulfur-cluster-free hydrogenase (Hmd) and F420-dependent methylenetetrahydromethanopterin dehydrogenase (Mtd), which both do not contain nickel [16,17]. The presence of nickel in hydrogenases from M. marburgensis was discovered by Graf and Thauer [18]. F420 -nonreducing [Ni-Fe]-hydrogenase (MvhADG). The cytoplasmic enzyme is composed of three different subunits of which MvhA is the subunit harboring the [NiFe] center and MvhD and G are iron–sulfur proteins. The hydrogenase is involved in CoM-S-S-CoB reduction with H2. Electron transfer between MvhADG and heterodisulfide reductase HdrABC probably proceeds between MvhD and the subunit HdrA [19]. Met. Ions Life Sci. 2, 323–356 (2007)

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O H S

–O S 3

CH 3

+

N

HS

H

H

Coenzyme B

Methyl-coenzyme M

OPO 32–

∆G°′ = – 30kJ/mol

CH4 + –

O

H

S O 3S

S

–

CO2 CH3

N H

H

OPO 32–

Heterodisulfide

Figure 2. archaea.

Reaction catalyzed by methyl-coenzyme M reductase from methanogenic

Energy converting [Ni-Fe]-hydrogenase isoenzyme I (EhaA-T) and isoenzyme II (EhbA-Q). Several subunits of the two enzymes show high sequence similarity to the energy-conserving NADH:quinone oxidoreductase (complex I) from various organisms. EhaA-T is composed of 20 different subunits of which EhaO harbors the active site [NiFe] center. EhbA-Q is composed of 17 different subunits of which EhbN is the subunit harboring the [NiFe] center [20]. EhaA-T is involved in CO2 reduction to methane with H2 at the oxidation level of CO2 and EhbA-Q has a role in autotrophic CO2 fixation. The two membrane-associated enzymes catalyze the proton or sodium motive-force-driven reduction of ferredoxin with H2 [5]. The presence of energy converting hydrogenases in methanogenic archaea was discovered by Künkel et al. [21]. Carbon monoxide dehydrogenase (CdhA). The cytoplasmic enzyme, which harbors a nickel–iron–sulfur cluster [22–24] is involved in autotrophic CO2 fixation [25,26]. It catalyzes the reduction of CO2 to CO with reduced ferredoxin. The first evidence that carbon monoxide dehydrogenase in anaerobic microorganisms contains nickel was published by Diekert et al. [27] and Drake et al. [28]. Acetyl-CoA synthase/decarbonylase (CdhBCDE). The cytoplasmic enzyme is composed of four different subunits of which CdhC harbors a binuclear Ni–Ni center associated with an iron–sulfur cluster [29–31] and CdhDE is an iron–sulfur corrinoid protein comples. The enzyme is involved in autotrophic CO2 fixation [25,26]. It catalyzes the reversible synthesis of acetyl-CoA from CO, N5-methyltetrahydromethanopterin, and CoA [32]. The first indications that the Met. Ions Life Sci. 2, 323–356 (2007)


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active site of acetyl-CoA synthase/decarbonylase contains nickel came from Shin and Lindahl [33,34]. The presence of two nickel was demonstrated only recently [29,35]. For the synthesis of the eight different nickel enzymes in M. marburgensis approximately 1 µmol nickel per g cells (dried mass) are required under nickel sufficient conditions [1]. Most of the nickel is incorporated into the nickel corphin coenzyme F430 [36], which is the prosthetic group of the two methyl-coenzyme M reductase isoenzymes. The two isoenzymes together are present in M. marburgensis in concentrations of up to 10% of the soluble cell proteins [4]. Nickel uptake by growing cells of M. marburgensis is reduced to 0.15 µmol/g when the concentration of nickel in the growth medium is growth rate limiting [1], but even under these conditions the nickel requirement is much higher than that of Escherichia coli. The nickel requirement of E. coli is mainly for the synthesis of its [Ni-Fe]-hydrogenases and is so low that it is generally covered by the nickel contaminations in the growth medium, which are in the order of 50 nM [1]. This is why the nickel requirement of E. coli and the presence of nickel in its hydrogenases were, for a long time, overlooked. The nickel enzymes found in M. marburgensis are present in most other methanogenic archaea (Methanobacteriales, Methanococcales, Methanopyrales, Methanomicrobiales, and Methanosarcinales) and most also occur in other archaea and in bacteria. Of the nickel enzymes only methyl-coenzyme M reductase is characteristic of all methanogenic archaea and of the phylogenetically closely related methanotrophic archaea [37]. These microorganisms can therefore be identified via their genes for this nickel enzyme. The genes coding for the three methyl-coenzyme M reductase subunits are generally coded in a transcription unit mcrBDCGA and/or mrtBDGA mrtC. The function of McrC and McrD, which are synthesized, but do not co-purify with methyl-coenzyme M reductase, is not known. In some methanogens and methanotrophic archaea both the mrtC and the mrtD genes are not located together with the mrtBGA genes. The phylogenetic tree deduced from the primary structure of the α subunit (McrA) of methyl-coenzyme M reductases I and II mirrors the one deduced from the 16 S RNA sequences of these microorganisms quite well [38–43]. As methyl-coenzyme M reductase, also coenzyme F430 is found only in methanogenic and methanotrophic archaea. The presence of the nickel corphin in a sample can be determined at very high sensitivity via MALDI-TOF mass spectrometry [42]. Many methanogens contain only one methyl-coenzyme M reductase. The presence of two isoenzymes appears to be restricted to some members of the orders of Methanobacteriales and Methanococcales. In Methanosarcina species, Methanococcus maripaludis, and Methanopyrus kandleri, which contain only one methyl-coenzyme M reductase, the primary structure of the enzyme is most similar to that of isoenzyme I (Mcr) of M. marburgensis. In Methanosphaera stadtmanae, which belongs to the order of Methanobacteriales and also contains Met. Ions Life Sci. 2, 323–356 (2007)

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only one methyl-coenzyme M reductase, the enzyme is most closely related to isoenzyme II (Mrt) of M. marburgensis [6]. In the following section first the structure and properties of the free nickel corphin cofactor F430 will be described and then those of the methyl-coenzyme M holoenzyme from methanogenic and methanotrophic archaea.

2.

STRUCTURE AND PROPERTIES OF COENZYME F430

F430 is released from methyl-coenzyme M reductase or whole cells of methanogenic archaea upon denaturation with perchloric acid or trichloroacetic acid. In the released cofactor, nickel is in the high-spin Ni(II) oxidation state. Due to its five carboxyl groups (Figure 1) the cofactor is extremely soluble in water, a property which is lost after conversion of F430 into the pentamethyl ester (F430M) [44] or pentaalkylamide [45]. These Ni(II)F430 derivatives are soluble in nonpolar, noncoordinating solvents, in which they are diamagnetic. The structures and properties of F430 and of F430M have been reviewed by one of us in Volume 29 of Metal Ions in Biological Systems [46]. Therefore, mainly the literature published after 1993 is discussed here. The discovery and biosynthesis of coenzyme F430 [47] and of other natural hydroporphyrins have been reviewed [48].

2.1. Structure of F430, of F430 Isolation Artifacts, and of a Natural F430 Derivative The constitution and configuration of F430M in the Ni(II) oxidation state (Figure 1) was established in a series of biosynthetic incorporation experiments with selectively labeled 13C-δ -aminolevulinic acid, spectroscopic studies, and by crystal structure elucidation [44,49–54]. The structure exhibits several unique elements. The π chromophore extends over only three of the four nitrogens and makes F430 the most extensively reduced tetrapyrrole found in nature. F430 is a tetrahydrocorphin, corphin being the name proposed by Eschenmoser [55] for this class of tetrapyrroles, in which the carbon framework of a porphin is combined with the linear chromophore typical for corrins. Two additional rings are found in F430, the lactam ring fused to ring B and the six-membered carbocycle formed through intramolecular acylation of meso position C-15 by the propionic acid side chain at ring D. Formation of this ring is the last step in the biosynthesis of F430, the direct precursor being 15,173-seco-F430 [56]. The fact that this annelation brings the carbonyl function into conjugation with the π chromophore has a profound influence on the redox properties of F430 [46]. For example, it has been calculated that the carbonyl substituent at the 15-meso position electronically stabilizes the Ni(I) state of the cofactor [57]. Met. Ions Life Sci. 2, 323–356 (2007)


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F430 is a labile compound, which slowly epimerizes to 13-epi-F430 and 12,13-diepi-F430 at room temperature and slowly oxidizes at room temperature to 12,13-didehydro-F430 in the presence of O2 [58,59]. Consistent with experiment, molecular mechanics calculations predict that Ni(II)-diepi-F430 is thermodynamically more stable than native Ni(II)F430 [60,61], but that, when bound in methyl-coenzyme M reductase, Ni(II)F430 rather than 12,13-diepi-Ni(II)F430 has the lower energy conformation [62]. Besides the isolation artifacts small amounts of 19,20-didehydroF430 with an absorption band at 339 nm are formed during purification of F430 [63]. F430, as shown in Figure 1, has a molecular mass of 905 Da and this is the mass of the nickel corphin cofactor in all methanogenic archaea as determined by MALDI-TOF mass spectroscopy. It was therefore surprising to find that the cofactor associated with one of the methyl-coenzyme M reductases isolated from methanotrophic archaea has a molecular mass of 951 Da [42]. In not yet published experiments it was established that the 951 Da cofactor is 172-methylthio-F430 (S. Mayr and B. Jaun, 2005, unpublished). Interestingly, the methylthio group is in the α position of the carbonyl group in the carbocyclic ring and is therefore predicted to have an effect on the redox properties of F430.

2.2.

Properties of Ni(II)F430 and Nonplanarity

Ni(II)F430M in dry dichloromethane, chloroform or acetonitrile is in a low-spin state. In this state the nickel in F430 is square planar tetracoordinated, but has the pronounced tendency to bind additional ligands in the axial direction such as chloride with log K1 5.4 and log K2 3.7 and imidazole with log K1 2.7 and log K2 2.2. Upon addition of the first axial ligand the electronic configuration changes from low-spin to high-spin [46]. The UV-visible spectrum of low spin F430M has an absorbance maximum at 442 nm (ε 21 mM1 cm1). Upon formation of five-coordinate complexes the absorption maximum shifts to shorter wavelength and the band gains intensity. Transition from five- to six-coordinate forms leads to a shift to longer wavelengths and a further increase in extinction. Low spin Ni(II)F430 is nonplanar due to the fact that low-spin Ni(II) is too small for the macrocyclic ligand system, forcing the pyrrole rings out of plane. This is even more so in Ni(III)F430 (see Section 2.4). The size of the nickel ion increases in the order Ni(III) low-spin Ni(II) high-spin Ni(II) Ni(I). With increasing size the tension in the macrocycle of F430 decreases and the ring becomes planar, as in high-spin Ni(II)F430 and Ni(I)F430 [46]. Different conformations of the ring system and its consequences have also been studied in Ni(II)F430 model compounds [62,64–69]. For an extensive discussion of the relationship between the structure of F430 and its redox properties the reader is referred to the reviews by Jaun [70] and by Ghosh et al. [57] on the subject. The redox chemistry of nickel complexes in aqueous solutions has recently been reviewed by Zilbermann et al. [71]. Met. Ions Life Sci. 2, 323–356 (2007)

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2.3. Reduction of Ni(II)F430 to Ni(I)F430 and the Properties of Ni(I)F430 Ni(II)F430M in acetonitrile [72] and Ni(II)F430 in H2O [73] can be reversibly reduced to the Ni(I) oxidation state at a redox potential (Eo) of approximately 600 mV compared with the normal hydrogen electrode (NHE). One F/mol are taken up showing that a single electron is transferred in the process [72]. The redox potential is comparable to that needed to reduce cob(II)alamin to cob(I)alamin. It is lower than that of the H/H2 couple (Eo 414 mV). Despite this fact, reduction of Ni(II)F430 to Ni(I)F430 in water is possible [73] since at pH values above 7 Ni(I)F430 reacts with protons only slowly to give H2 and Ni(II)F430. In the presence of O2 Ni(I)F430 is almost instantaneously oxidized to Ni(II)F430. Nickel in Ni(I)F430M has been found to be tetracoordinated under all experimental conditions tested. But other Ni(I) hydroporphyrins have been shown to bind a single axial ligand [67]. As indicated above in Section 2.2, Ni(II)F430M can be tetra-, penta- and hexa-coordinated. Since the tendency of Ni(I)F430 and Ni(II)F430 to bind axial ligands is very different, it can be predicted that the redox potential of the Ni(II)F430 / Ni(I)F430 couple will be strongly affected by the presence and nature of ligands, that can axially be coordinated to Ni(II)F430 and/or Ni(I)F430. The UV-visible spectra of Ni(I)F430M and of Ni(I)F430 show an intense absorbance maximum at 383 nm (ε 34 mM1 cm1; MeCN) and a weak one at 760 nm. The EPR spectrum of Ni(I)F430 is of nearly axial type with g⊥ g储. A partially resolved hyperfine splitting on the g⊥ line is assigned to the coupling with the four nitrogen nuclei of the equatorial macrocycle. The best fit of the spectrum is obtained with the parameters gx 2.065, gy 2.074 and gz 2.250 and a single isotropic aN value of 0.95 mT. The strong anisotropy, the order of the g values and the equatorial hyperfine coupling constants are consistent with a species of effective spin S 1兾2 with its spin population residing predominantly in the d x2y2 -type orbital of the central nickel ion [72]. Also at Q-band, the g tensor of Ni(I)F430 in H2O appears axial and ENDOR spectra of Ni(I)F430 in H2O versus D2O solvent show no evidence for strongly coupled, solvent-exchangeable hydrogen, indicating that there is no water axially coordinated to Ni(I) in contrast to the Ni(II)F430 state [74,75]. The nickel center in Ni(I)F430 is truly Ni(I): it carries approximately 82% of the total molecular unpaired spin [61]. The observation that Ni(I)F430M can only be further reduced at potentials below 1.6 V indicates that reduction of F430 to the Ni(0) valence state is not possible under physiological conditions [46]. Recently it has been reported that the reduction of Ni(II) F430 with Ti(III) is a three-electron rather than a one-electron process, indicating that besides Ni, a double bond of the hydrocorphinoid macrocycle is reduced as well [76]. The reduction product had the same UV-visible and EPR spectrum as Ni(I)F430M published to be generated from Ni(II)F430M in a one-electron reduction [72]. To resolve this Met. Ions Life Sci. 2, 323–356 (2007)


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contradiction, Piskorsky and Jaun [77] have reinvestigated the stoichiometry of the reduction of F430M by three independent methods. Spectroelectrochemistry showed clean reduction to a single product that exhibits the UV-vis spectrum described for Ni(I)F430M and Ni(I)F430 generated by reduction of Ni(II)F430 with Ti(III). In three bulk electrolysis experiments, 0.96 ± 0.1 F/mol was required to generate the reduced species. Reduction with decamethylcobaltocene in tetrahydrofuran consumed 1 mol the cobaltocene per mol F430M, and the stoichiometry of the reoxidation of the reduced form with the two-electron oxidant methylene blue was 0.46 ± 0.05 mol methylene blue per mol reduced F430M. These experiments demonstrate that the reduction of F430M is a one-electron process and therefore inconsistent with a reduction of the macrocycle chromophore. Craft et al. [78] employed DFT calculations to evaluate possible reduced F430 models: a one-electron reduced Ni(I)F430 model and a three-electron reduced Ni(I)F430-red model (possessing a reduced hydrocorphin ligand) on the basis of excited-state spectra and of published EPR/ENDOR parameters. While calculations on both models yield spectroscopic parameters that are consistent with most experimental data, overall better agreement was achieved using the Ni(I)F430 model, particularly with respect to electronic absorption and ENDOR. The experimentally validated bonding descriptions generated show that in Ni(II)F430 the occupied Ni 3d orbitals are too low in energy to significantly perturb the dominant electronic transition involving the π and π* frontier MOs of the macrocycle (i.e., the HOMO→LUMO transition). Upon one-electron reduction of the Ni(II) ion, the occupied Ni 3d orbitals are raised in energy, shifting between the HOMO and the LUMO of the oxidized cofactor. These ground-state changes have a dramatic effect on the excited-state structure, causing a blue shift of the dominant π→π* transition and the appearance of numerous Ni 3d→hydrocorphin π* charge-transfer features in the vis/near-IR region [78]. Ni(I)F430 is nucleophilic. It reacts with protons to H2 and with alkylhalides to alkanes, but not with methyl-coenzyme M [46]. Coenzyme F430 can be used to catalyze the reductive dehalogenation of chlorinated or brominated hydrocarbons [79–81]. Several F430 model compounds have been shown to have the same property [82–86]. The reaction of alkylhalides with Ni(I)F430 can mechanistically explain why 2-bromoethanesulfonate and 3-bromopropanesulfonate are suicide inhibitors of active methyl-coenzyme M reductase [87].

2.4. Oxidation of Ni(II)F430M to Ni(III)F430M and the Properties of Ni(III)F430M Ni(II)F430M in acetonitrile can be reversibly oxidized at a redox potential of 1.6 V compared with NHE with 1 F/mol being transferred. Ni(III)F430M is not stable in the presence of water (Eo 0.81 V) or of other ligands with redox potentials distinctly lower than 1.6 V. Met. Ions Life Sci. 2, 323–356 (2007)

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The UV-visible spectrum of Ni(III)F430 shows an absorption maximum at 360 nm (ε 16 mM1 cm1; MeCN) and the EPR spectrum consists of a strongly anisotropic signal of the axial type (g⊥ 2.211 and g储 2.020) with barely resolved fine structure on the g储 line and no resolved structure on the g⊥ part. This type of spectrum is typical for an S 1兾2 species with the unpaired electron in a nickel dz2 orbital, the configuration which is expected for a Ni(III) ion in an ‘elongated’ tetragonal field. The fine structure of the g储 line is due to the coupling with the nitrogen nuclei of two acetonitrile solvent molecules coordinated in the axial position [46]. Ni(III)F430 is probably always hexa-coordinated. The very positive redox potential of the Ni(III)F430M/Ni(II)F430M couple in acetonitrile appears to exclude Ni(III)F430 as intermediate in the enzymatic process. However, stabilization of the Ni(III) oxidation state by axial ligands stronger than acetonitrile is expected to reduce the oxidation potential towards more cathodic potentials. Thus, in cyclic voltammograms, dramatic cathodic shifts of peak potentials of up to 0.5 V were indeed observed in the presence of potential ligands such as imidazole. However, it was not possible to deduce thermodynamic data from these experiments because the reversibility of the oxidation process was partially lost in the presence of these ligands [46]. Based on the results of DFT calculations, Wondimagegn and Ghosh [88] predicted that Ni(III)F430 with strong axial ligands (or high spin Ni(II)F430 axially coordinated to a radical) would be a reasonable intermediate in the catalytic cycle of methyl-coenzyme reductase. Using density functional theory in 2001 Ghosh and Steene [90] carried out a quantum chemical survey of high-valent transition metal porphyrins and related compounds amongst them Ni(III)F430. They discussed whether the molecules feature ‘true’ high-valent metal centers, or whether the ligands are oxidized instead, i.e., are non-innocent, and come to the conclusion that the electronic structures fall somewhere along the continuum between these scenarios [89,90].

2.5. Methyl-Ni(II)F430 and Methyl-Ni(III)F430 As indicated in Section 2.3, Ni(I)F430 does not react with methyl-coenzyme M. However, with more reactive electrophiles, such as methyl iodide ⯝ methyl triflate methyl dialkyl sulfonium hexafluorophosphate methyl tosylate, Ni(I)F430M reacts to give methane and Ni(II)F430M, the fourth hydrogen of methane being introduced as a proton rather than as a hydrogen atom [91,92]. If methyl iodide was allowed to react with Ni(I)F430M in a 2:1 stoichiometric ratio at low temperature, the color of the solution changed within seconds from green Ni(I)F430M to brown-orange without methane being formed. If acid was added, accompanied by a color change to that of Ni(II)F430M, more than 80% of the theoretical amount of methane was generated. Addition of deuterated acidic acid gave over 85% CH3D [91]. This experiment proves that an intermediate is formed, which can be dissociated to Ni(II)F430M and methane by protonation. Its properties are consistent Met. Ions Life Sci. 2, 323–356 (2007)


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with those expected for a methyl-Ni(II) derivative with an axial nickel–carbon bond, which was verified for CD3-Ni(II)F430M by NMR spectroscopy [93]. The dissociation energy for the Ni(II)–C bond was estimated to be of the order of 80 kJ/mol [94]. As intermediate in methyl-Ni(II)F430M formation from Ni(I) and methyl iodide methyl-Ni(III)F430M is discussed. However, all attempts to demonstrate the existence of this compound have failed until now, although theoretical modeling is consistent with methyl-Ni(III)F430 as intermediate in the catalytic mechanism of methyl-coenzyme M reductase [88]. Based on the peak potential of the irreversible oxidation of methyl-Ni(II)tetramethylcyclam, a model compound for methyl-Ni(II)F430, the redox potential of the methyl-Ni(III)F430 /methyl-Ni(II)F430 couple is estimated to be 450 mV vs NHE (T. Vogelsang, Diploma Thesis, ETH Zürich, 1995, unpublished).

2.6. Model Reaction for Methyl-Coenzyme M Reduction with Coenzyme B to Methane The formation of a sulfuranyl radical intermediate followed by methyl transfer to the Ni(I) center of F430 and generation of the disulfide has been proposed as a possible mechanism for the formation of methane catalyzed by methylcoenzyme M reductase in methanogenic archaea [95–97]. In order to test this hypothesis, a sterically shielded, bifunctional model substrate that contained a methyl thioether and a sulfhydryl functional group, which could form a fivemembered cyclic sulfuranyl radical according to the postulated mechanism, was synthesized. The corresponding thiolate reacted with Ni(II) salts to give a diamagnetic, square planar Ni(II) dithiolate complex, which was characterized by X-ray diffraction. Upon irradiation of this complex with light of λ 300 nm, methane and the cyclic disulfide were formed, whereas irradiation of the thiolate in the absence of nickel gave only traces of methane and no cyclic disulfide. The observed products are consistent with the postulated mechanism via a sulfuranyl radical, and the role of light is interpreted as the formation of a Ni(I)/thiyl radical pair upon excitation of a charge-transfer band of the Ni(II) dithiolate. In the presence of a large excess of thiolate, the diamagnetic complex was transformed into a paramagnetic, five- or six-coordinate complex that proved to be more active in the generation of both methane and the cyclic disulfide than the square planar diamagnetic dithiolate [98]. Although this model reaction yields methane and a disulfide from a methyl thioether and a thiolate, its mechanism most probably differs from that of enzyme-catalyzed methane formation. Thus the model reaction starts with a light-driven reduction of Ni(II) with a thiol to Ni(I) and a thiyl radical, whereas the methyl-coenzyme M reductase reaction almost certainly starts with the oxidation of Ni(I) with methyl-coenzyme M (see Section 4.3). Met. Ions Life Sci. 2, 323–356 (2007)

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For completeness, two other reports on models for the in vivo activity of coenzyme F430 are mentioned here: Drain et al. [99] reported that methyl-coenzyme M is converted to methane using the nickel macrocyclic complex [1,4,7,10,13pentaazacyclohexadecane-14,16-dionato] Ni(II) in aqueous solution. During the process also O2 was formed and the process was not stimulated by reductants. Reinvestigation of this reaction revealed, however, that it was catalyzed by an impurity present in the technical grade tetraethylenepentamine rather than by the nickel complex [100]. The second report is by Zilbermann et al. [101] who described that methyl-coenzyme M is reduced by monovalent macrocyclic nickel complexes at pH 9.4 to methane in a 10% yield involving methyl free radicals as intermediate. Reduction of the Ni(II) complex to the Ni(I) complex was achieved by irradiating He saturated solutions in 0.01 M sodium formate with ionizing radiation. The preliminary communication was not followed up by a more detailed investigation.

2.7. Models for Coenzyme M in the Active Site of MethylCoenzyme M Reductase The crystal structure of methyl-coenzyme M reductase (see Section 3.1) revealed that the Ni center in the prosthetic group of coenzyme F430 is penta- or hexacoordinated with the carboxamide group of a glutamine residue occupying the axial coordination site on the α-side of the macrocycle. To obtain diastereoselectively coordinated complexes for mechanistic and spectroscopic studies of the free coenzyme in solution as models for F430 bound within the active site, Bauer and Jaun [102,103] prepared partial-synthetic derivatives of coenzyme F430 that have a coordinating group attached via a tripeptide linker to one of the propanoic acid side chains. Suitable linkers were evaluated based on molecular mechanics and NMR solution structures and the most promising linker, Pro-Pro-His(π-Me)OMe was coupled via its N-terminus to the free carboxylate side chain at C(3) of F430 tetramethyl ester. The UV/visible and NMR spectra in CH2Cl2 /3,3,3-trifluoroethanol 6:1 show that the new derivative, the Ni(II) (33-dehydroxy-83,122,133,182tetra-O-methyl-F430-33-yl)-L-prolyl-L-prolyl-N-π-methyl-L- histidine methyl ester is an intramolecular, pentacoordinate, paramagnetic complex. In the same solvent system, the parent 33,83,122,133,182-penta-O-methyl-F430 ester is four coordinate and diamagnetic even in the presence of equimolar 1-H-imidazole. Protonation of the axially coordinating histidine residue gave the diamagnetic tetracoordinate base-off form, which allowed to establish its constitution by NMR [102,103]. Likewise, tetraazamacrocyclic nickel(II) complexes with aliphatic methylthioand methoxy-substituted pendant chains were shown to have a coordination number of 5 and 4, respectively. Cyclic voltammetry of these complexes in acetonitrile revealed that Ni(II) is reversibly reduced to Ni(I) between 0.64 and 0.77 V compared with the standard calomel electrode (SCE), the potential being influenced by the nature and number of the pendant chains. At more negative potentials, the Met. Ions Life Sci. 2, 323–356 (2007)


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thioether was cleaved, whereby a thiol was formed; the thiol was then oxidized at ca. 0.8 V compared with SCE, when a glassy carbon electrode was used. No cleavage of the ether bond was observed under similar conditions [104,105].

3.

MOLECULAR PROPERTIES OF METHYL-COENZYME M REDUCTASE

A detailed review on the structure and properties of methyl-coenzyme M reductase has been published by one of us in 1998 [4]. Therefore, only results obtained after 1998 will be discussed here in greater detail. Recent reviews on methylcoenzyme M reductase are by Ermler [106] and Shima and Thauer [37]. Most of what is known about methyl-coenzyme M reductase comes from work on the isoenzyme I (McrABG) from Methanothermobacter marburgensis, which has a growth temperature optimum of 65C. Isoenzyme I from this thermophilic methanogen is in the following referred to as MCR. Available evidence indicates that most of the structural properties of this enzyme are shared by isoenzyme II from M. marburgensis and by methyl-coenzyme M reductase from other methanogenic archaea. Where this is not the case, this will be indicated. All methyl-coenzyme M reductases analyzed to date have a molecular mass of approximately 300 kDA, are composed of three different subunits in an α2β2γ2 arrangement, and contain two molecules of the nickel corphin coenzyme F430 tightly, but noncovalently bound. The coenzyme has to be in the Ni(I) oxidation state for the enzyme to be active. Active MCR is referred to as MCRred1 and its EPR signal as the MCRred1 signal. Under oxic conditions MCRred1 is almost instantaneously inactivated with the concomitant loss of the Ni(I)-based EPR signal. This inactive form is referred to as MCRred1-silent. In vivo the EPR-silent forms can be converted back to the active form in an enzyme-catalyzed, ATP-dependent complex reduction process. The reactivation system involved has, until now, been only partially characterized [4]. MCRred1 can be converted to other EPR active forms which are distinguished via differences in their EPR spectra: MCRred1/2; MCRox1; MCRox2; MCRox3, MCRBPS They are generated from MCRred1 in vitro in the following reactions: MCRred1 HS-CoM HS-CoB MCRred1/2 polysulfide MCRred1/2 sulfite MCRred1/2 O2 MCRred1 3-bromopropane sulfonate MCRred1 O2 MCRox1 (slowly) MCRox1 Ti(III)

L → → → → → → →

MCRred1/2 MCRox1 MCRox2 MCRox3 MCRBPS MCRred1-silent MCRox1-silent MCRred1

(1) (2) (3) (4) (5) (6) (7) (8)

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The MCRox- and MCRsilent forms are enzymatically inactive as is MCRBPS. In the presence of coenzyme M (HS-CoM), which is a substrate analog of methyl-coenzyme M, the enzymatic activity of MCRred1 is reversibly inhibited. MCRred1/red2 and MCRox1 are also observed in vivo [107]. They are generated from MCRred1 in intact cells when these are gassed with 100% H2 and 80%N2 /20%CO2, respectively. Methyl-coenzyme M reductase isolated from M. marburgensis cells after harvest without prior gassing of the cells with 100% H2 is EPR silent and referred to as MCRsilent [4]. In the different MCR states the nickel in F430 is in different oxidation and coordination states as indicated by their crystal structure and/or EPR spectra, MCD spectra, X-ray absorption spectra, and UV-visible spectra.

3.1. Crystal Structures of MCRox1-silent and MCRsilent Until now crystal structures of MCR are only available for inactive, EPR-silent forms, in which the nickel of F430 is in the Ni(II) oxidation state [108–110] (for a review see [111]). The structure of the inactive enzyme, the best structure resolved to 1.16 Å, is characterized by a series of α helices arranged in a compact form with an ellipsoidal shape of about 120 A 85 A 80 A or 120 85 80 A exponent 3 (A Angstrom). There are two identical F430 binding sites, roughly 50 Å apart. Each F430 is buried deeply within the protein complex and accessible from the proteins surface only via a 50-Å-long channel, which at its narrowest part is only 6 Å in diameter. The channel and the coenzyme binding sites are formed mainly by hydrophobic residues of subunits α, α’, β and γ and α’, α, β’ and γ’, respectively, i.e., both α subunits are involved in forming both active sites. Thus, whereas the corphin ligand system of F430 is tightly attached to one α subunit, the lower axial ligand to nickel is contributed by a glutamine residue of the second α subunit. Via this architecture the two active sites are structurally interlinked despite the fact that a distance of nearly 50 Å separates them. This is quite unique. F430 is bound at the bottom of the 50-Å-long channel such that the tetrapyrrole plane of F430 points with its front face (β -face, reduced pyrrole rings A, B, C, and D clockwise) (Figure 1) roughly towards the mouth of the channel, whereas its rear face (α face) points to the bottom of the channel. Thus only the front face is accessible to the substrates. The tetrapyrrole ring of F430 is bound in a rather flat conformation to the enzyme as predicted for free Ni(II)F430 in the hexacoordinated state. The Ni atom sits almost exactly in the tetrapyrrole plane and is coordinated to six ligands arranged in a nearly optimal octahedral configuration. Four of the ligands are provided by the four equatorially located nitrogen atoms of the tetrapyrrole ring. As the fifth ligand the side chain oxygen of Gln α147 protrudes from a long loop to the rear face of F430 and approaches the Ni(II) atom to 2.3 Å. Gln α147 is embedded in the protein matrix such that it will probably not be able to move away (or only after a mayor conformational change of the Met. Ions Life Sci. 2, 323–356 (2007)


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Figure 3. Binding of coenzyme M and of coenzyme B in the active site in MCRox1silent (A) and of the heterodisulfide CoM-S-S-CoB in the active site of MCRsilent (B). For details see Section 3.1.

matrix) when F430 is reduced from the Ni(II) to the Ni(I) oxidation state, in which the Ni(I) is preferentially tetra-coordinated [46,74]. It has been suggested that by the enforced axial fifth ligand the nucleophilicity of the Ni(I) in F430 is increased [4]. It can also be envisaged that upon reduction of the Ni(II) to the Ni(I) state the upper axial ligand of the hexa-coordinated Ni(II) is actively released. In turn, an enforced upper axial ligand to Ni(I) such as in the MCRred2 state (see Section 3.3) could force the lower ligand to move away. MCRox1-silent contains coenzyme M and coenzyme B bound (Figure 3A). Coenzyme M is coordinated via its thiol sulfur to Ni(II) of F430 (2.42 Å distance). Whether the thiol group is coordinated as such or only after dissociation to the thiolate is not known. The thiol/thiolate group binds axially to the Ni(II) and interacts with the hydroxyl group of Tyr α333 and Tyr β367 and a water molecule that bridges coenzyme M and coenzyme B. Coenzyme B, in its elongated conformation, fits accurately into the most narrow segment of the channel formed by residues of the subunits α, α’, and β. The thiol group of coenzyme B is positioned at a distance of 8.7 Å from the nickel of F430 and 6.2 Å from the thiol sulfur of coenzyme M [4]. MCRsilent contains the hetrodisulfide CoM-S-S-CoB bound (Figure 3B). A superposition of the structures reveals that the reduced coenzyme B in Met. Ions Life Sci. 2, 323–356 (2007)

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MCRox1-silent and the coenzyme B moiety of the heterodisulfide in MCRsilent align perfectly, except that the sulfur is turned slightly towards coenzyme M. In contrast to coenzyme B, coenzyme M has moved more than 4 Å away from its position in the MCRox1-silent state. The thiol group is shifted perpendicular and the sulfonate group parallel to the tetrapyrrole plane of F430, resulting in a 90 rotation of coenzyme M. In this position one oxygen atom of the sulfonate is axially coordinated to the nickel and contacts the hydroxyl group of Tyr α333. The distance between nickel and the oxygen is 2.1 Å. A second oxygen atom of the sulfonate group is hydrogen bonded to the lactam ring of F430 and to the hydroxyl group of Tyr β367 and a third one to a water molecule located at the former binding site of the sulfonate group [4]. Near the active site are five modified amino acids: a thioglycine, a N-methylhistidine, a S-methyl cysteine, a (5S)-5-methyl arginine and a (2S)-2-methyl glutamine [108–111]. Labeling studies have shown that the methyl groups are biosynthetically derived from the methyl group of methionine and not from the methyl group of methyl-coenzyme M [112]. The five modified amino acids, all in the α subunit, are highly conserved. Only in MCR from Methanosarcina the respective glutamine is not methylated [109]. The thioglycine forms a thiopeptide bond, which might be susceptible to reduction induced trans–cis isomerization (DFT calculations by B. Jaun, unpublished; analogy: light-induced cis–trans isomerization [113]) and could therefore play a key role in coupling of the two active sites. Until now methyl-coenzyme M reductase is the only protein known to harbor a thiopeptide group and all methyl-coenzyme M reductases analyzed in this respect have been shown to contain the unusual bond. Besides McrA from M. marburgensis the analyses have included MrtA from M. marburgensis, from Methanococcus voltae, from Methanopyrus kandleri, and from Methanoculleus thermophilus (J. Kahnt and R. Thauer, 2006, unpublished) and McrA from Methanosarcina barkeri [109]. The analyses were performed essentially as described by Selmer et al. [112].

3.2.

MCRred1 with Penta-Coordinated Ni(I)F430

MCRred1 is the only form of MCR that is enzymatically active. Fully active MCR has a greenish color [114,115]. Its UV-visible spectrum shows an intense band at 383 nm and a weaker one centered around 760 nm und is almost identical to that of free Ni(I)F430 or Ni(I)F430M. Also, the EPR signal of MCRred1 (gz 2.2500; gy 2.0710; gx 2.0605) is very similar to that exhibited by Ni(I)F430M (gz 2.250; gy 2.074; gx 2.065). The findings indicate that in MCRred1 F430 is in the Ni(I) oxidation state. In the fully active enzyme both F430 are in the Ni(I) state as indicated by a spin concentration of above 0.9 (spin per Ni) determined by double integration of the EPR signals. There is a positive correlation between spin concentration and enzyme activity. The MCRred1 EPR signal is Met. Ions Life Sci. 2, 323–356 (2007)


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almost immediately quenched by O2 and very rapidly by chloroform [116] and 2-bromoethanesulfonate [87]. MCRred1 is very labile, even under strictly anoxic conditions. It is stabilized considerably, however, in the presence of coenzyme M, which is why MCRred1 is purified in the presence of this substrate analog and competitive inhibitor [87]. Upon addition of coenzyme M to MCRred1, its EPR signal changes only slightly. MCRred1 in the presence of coenzyme M is referred to as MCRred1c [117,118]. X-ray absorption spectroscopy revealed that the nickel in MCRred1c is coordinated by five N/O ligands (most probably four nitrogen ligands from F430 and one lower oxygen ligand from glutamine α147) and that the coordination state does not change when MCRred1 is incubated in the presence of methyl-coenzyme M and/or coenzyme B [118]. This finding indicates that stabilization of MCRred1 by coenzyme M is not via binding of the competitive inhibitor to the active site nickel. On the basis of XANES and EXAFS data Tang et al. [76] and Singh et al. [119] have reported that the Ni(I) in MCRred1 is best described as being hexa-coordinate with six N/O ligands. The conflicting result can be explained assuming that the samples analyzed contained significant amounts of MCRsilent, in which the Ni(II) is hexa-coordinated by 4 N and 2 O (see Section 3.1, Figure 3B). Indeed the spin concentrations of the samples were relatively low (between 57 and 63%) [76].

3.3. MCRred1/2 with Coenzyme M Coordinated with the Sulfur of Its Thiol Group to Ni(I) of F430 in One of the Two Active Sites When MCRred1 is incubated in the presence of coenzyme M (inhibitor) and coenzyme B (substrate) part of its axial red1 EPR signal (gz 2.2500; gy 2.0710; gx 2.0605) is converted to a novel rhombic, nickel-based EPR signal designated red2 signal (gz 2.2940; gy 2.2383; and gx 2.1790) and its color changes from greenish (absorption band at 383 nm) to brownish (broad absorption band centered at 416 nm with shoulders at 390 and 450 nm). Both, coenzyme B and coenzyme M, are required for the conversion which is fully reversible [115,117,120]. MCRred1 can be maximally converted to 50% into the MCRred2 state, the rest always remains in a state which is similar, but not identical to the MCRred1 state with gz 2.2745, gy 2.0820 and gx 2.0680. The maximally 50% conversion can best be explained assuming that induction of the red2 state in the one active site of the enzyme, via a conformational change, prevents its induction in the second site (half-of-the-sites reactivity) [121]. Methyl-coenzyme M reductase with one active site in a red1 like state and the other one in the red2 state is referred to as MCRred1/2 [121]. Equivalently, methyl-coenzyme M reductase with both active sites in the red1 state, can be referred to as MCRred1/1. The formation of MCRred1/2 can therefore be described Met. Ions Life Sci. 2, 323–356 (2007)

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by the following equation: MCRred1 / 1 HS-CoM HS-CoB MCRred1 / 2

(9)

The conversion is reversible and temperature-dependent. The equilibrium lies far to the right at temperatures above 20C and far to the left at temperatures near 0C. Thus simply by lowering or increasing the temperature MCRred1/1 can be converted to MCRred1/2 and vice versa [121]. Since the red2 state is in temperature-dependent equilibrium with the red1 state the red2 state must also contain Ni(I). In MCRred1/2 the reactivity of the Ni(I) in both active sites of methyl-coenzyme M reductase is altered. Thus, in the presence of polysulfide not only the red2 signal, but also the red1 signal of MCRred1/2 is converted to an axial, nickel-based EPR signal designated MCRox1 (gz 2.2310; gy 2.1667; and gx 2.1532) (see Section 3.5). The spin concentration of the ox1 signal is generally above 0.8 (spin per Ni) indicating that in both active sites the nickel is in the ox1 state (MCRred1/1 polysulfide → MCRox1/1). Whereas, under the same experimental conditions, MCRred1/1 is to 100% rendered EPR silent [122]. Via X-band EPR spectroscopy and Q-band HYSCORE spectroscopy and employing [2-33S]-coenzyme M it was shown that in the MCRred2 state coenzyme M coordinates via its thiol sulfur to Ni(I) of F430 [123, 124]. It is not yet known whether coordination is via the thiol group or the thiolate group. The fact that generation of the MCRred2 state is dependent on coenzyme B indicates that the coordination of coenzyme M to the Ni(I) is enforced by the binding of coenzyme B to the enzyme. Most probably a larger conformational change is induced upon binding of coenzyme B, as suggested by the change in the EPR signal from axial (red1) to rhombic (red2). From the spectroscopic data it is not yet clear whether the Ni(I) in the MCRred2 state is hexa- or penta-coordinated. In the latter case binding of the thiol group of coenzyme M to Ni(I) would have to be associated with the dissociation of the oxygen ligand from glutamine α’147, which is the α-axial ligand of penta-coordinated Ni(I) in MCRred1. Since the two active sites are structurally interlinked via the α-axial ligand, binding of the thiol group of coenzyme M to Ni(I) in one active site could affect the reactivity of the Ni(I) in the second active site.

3.4. MCRBPS with a Ni(III)-Alkyl Bond as One of Three Resonance Structures When MCRred1 is incubated in the presence of 3-bromopropanesulfonate (or 3-iodopropanesulfonate or 4-bromobutyrate) its EPR signal with gz 2.219, gy 2.0710 and gx 2.0605 is converted to an axial signal with gz 2.2500, gx,y 2.116 and its color changes from greenish (absorption band at 383 nm) to Met. Ions Life Sci. 2, 323–356 (2007)


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yellow (absorption band at 424 nm).The EPR signal generated by 3-bromopropane sulfonate is quenched upon exposure of MCR BPS to O2, but not by 2-bromoethanesulfonate [87,120]. EPR spectroscopy of MCR BPS revealed the presence of a nickel–alkyl bond [125]. The hyperfine interaction between the unpaired electron spin on the nickel of F430 and the isotopically enriched 13Cγ nuclear spin indicates a Ni-Cγ coordination with approximately 6% of the spin population on the nearly sp2-hybridized Cγ. Under the not unrealistic assumption that upon reaction of Ni(I) with 3-bromopropanesulfonate, bromide rather than a bromine atom is formed as the byproduct [125], the resulting species is on the oxidation level of alkyl-Ni(III)F430. However, the hyperfine data point to a nearly planar bonding geometry of Cγ, thus indicating that the actual bonding situation is nearer to the alkyl radicalNi(II)F430 or even alkyl cation-Ni(I)F430 resonance structures than to alkyl anionNi(III)F430.

3.5. MCRox1 with Coenzyme M Coordinated via the Sulfur of Its Thiol Group to Ni(III) of F430 as One of Two Resonance Structures MCRox1 is generated in vitro from MCRred1/2 in the presence of polysulfide and in vivo by gassing 80%H2 /20%CO2-grown cells with 80%N2 /20%CO2 [117]. Both conditions are assumed to be oxidizing. Polysulfide was shown to rapidly oxidize MCRred1 (Ni(I)F430) to MCRsilent(Ni(II)F430) [117]. MCRox1 is also formed from MCRox1-silent by γ -irradiation at low temperatures [126]. MCRox1 is relatively stable under oxic conditions, while MCRred1 is not. The nickel in MCRox1 is hexa-coordinated by 5 N/O and 1 S as revealed by X-ray absorption spectroscopy [118]. MCRox1 exhibits a visible spectrum with an absorption band centered at 415 nm. There is also a weak band at around 650 nm [115]. MCRox1 shows an axial EPR signal with gz 2.2310, gy 2.1667 and gx 2.1532 [122]. The signal is more similar to that of Ni(I)F430M (gz 2.250, gy 2.074 and gx 2.065) than to Ni(III)F430M (gz 2.020 and gx,y 2.211). EPR and ENDOR results have therefore been interpreted as indicating a Ni(I) oxidation state for MCRox1 [126,127]. The interpretation contradicted the finding that MCRred1 is formed from MCRox1 by reduction [114]. Raman data led to the proposal that MCRred1 and MCRox1 differ in the reduction state of the corphin ligand system rather than in the nickel oxidation state, the ligand system being more reduced in MCRred1 [76,119]. However this hypothesis is inconsistent with recently published redox titrations [77] and not supported by optical/MCD spectroscopic data and density function calculations [78]. Both, theoretical and experimental evidence was provided that the nickel in MCRox1 is best described using Ni(III) thiolate and Ni(II) thiyl radical resonance forms [88,115,128]. The latter Met. Ions Life Sci. 2, 323–356 (2007)

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interpretation is also most consistent with UV-visible absorption and XAS data [115,118]. Recently, a pulse EPR study of MCRox1 was performed providing a detailed description of the spin density and the coordination of the Ni in coenzyme F430. MCRred1 was purified from cells grown in a 61Ni-enriched medium and converted to MCRox1 either in H2O or in D2O with coenzyme M either deuterated in the β -position or with 33S in the thiol group. To obtain the magnetic parameters ENDOR and HYSCORE measurements were done at X- and Q-band, and CW EPR at X- and W-band. The hyperfine couplings of the β -protons of coenzyme M indicate that the nickel to β -proton distances in MCRox1 are very similar to those in Ni(II)MCRox1-silent (see crystal structure), and thus the position of coenzyme M relative to F430 is very similar in both species. The 33S and nickel EPR data prove a Ni–S coordination, with an unpaired spin population on the sulfur of 7 ± 3%. These results highlight the non-innocent nature of the sulfur ligand and explain the difficulties encountered with the assignment of the correct oxidation level based on the spectroscopic properties of the nickel. Assuming that MCRox1 is oxidized relative to the Ni(II) species, the complex is formally best described as a Ni(III)thiolate in resonance with a thiyl radical/high-spin Ni(II) complex [129]. Finally, two results remain to be explained. (i) Becker and Ragsdale [130] reported that MCRox1 can be generated in vivo by the addition of sodium sulfide to 80%H2 /20%CO2 grown cells and concluded from this finding that MCRox1 is formed under reducing rather than oxidizing conditions. This did not consider, however, that sodium sulfide solutions almost always contain some polysulfide and that only µ M concentrations of polysulfide are required for the oxidation of MCRred1/2 to MCRox1 [122]. (ii) Telser et al. [126] reported that MCRox1-silent is partially converted to MCRox1 by γ -irradiation at low temperatures. γ -Irradiation of metal proteins frozen in water generally leads to the reduction of metal centers by electrons kicked out from bulk water. However, in the case of MCRox1-silent also cryoxidation has to be considered. The crystal structure of MCRox1-silent shows a water molecule close to the Ni(II)thiolate group that could trigger oxidation of the thiolate ligand or Ni(II) center [115].

3.6. MCRox1-silent, MCRred1-silent and MCRsilent with Hexacoordinated Ni(II)F430 MCRox1-silent is slowly (within hours) generated from MCRox1 both under oxic and anoxic conditions. Under reducing conditions, e.g., in the presence of mercaptoethanol, the conversion appears to be somewhat more rapid. Conversely, MCRred1-silent is formed almost instantly when MCRred1 is exposed to O2

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and the conversion is slowed down under reducing conditions. MCRox1-silent, MCRred1-silent and MCRsilent have visible spectra similar to that of Ni(II)F430 when hexa-coordinated [46]. The spectrum of MCRsilent has a maximum at 420 nm with a shoulder near 445 nm. That of MCRred1-silent is only slightly different [115]. The MCD spectrum of the two EPR silent inactive forms indicates the presence of high spin Ni(II) [115]. The crystal structure of MCRox1-silent and MCRsilent shows a hexa-coordinated nickel with four nitrogen ligands from F430, one lower oxygen α-ligand from glutamine α147 and one upper ligand, which in the case of MCRox1-silent is a sulfur and in the case of MCRsilent an oxygen (Figures 3A and B) [108–111]. X-ray absorption spectra are consistent with the crystal structure. Best fits for the Ni EXAFS were obtained for MCRox1-silent assuming six-coordinate nickel with five nitrogen or oxygen ligands and one sulfur ligand and for MCRsilent assuming six-coordinate nickel with six nitrogen or oxygen ligands [118].

3.7. Methyl-Coenzyme M Reductase from Methanotrophic Archaea The microorganisms catalyzing the anaerobic oxidation of methane (AOM) have been shown to contain gene homologues of mcrBGA and of most of the other genes involved in CO2 reduction to methane in methanogenic archaea [41,131]. Methanotrophic archaea present in the microbial mats catalyzing AOM in the Black Sea were found to contain at least two different methyl-coenzyme M reductases designated Ni-protein I and Ni-protein II, which could be separated by anion exchange chromatography [42]. Ni-protein II contained normal F430 with a molecular mass of 905 Da, whereas Ni-protein I contained a modified F430 with a molecular mass of 951 Da. Ni-protein I was present in a concentration of 7% of the extracted soluble proteins and Ni-protein II in a concentration of up to 3%. The N-terminal amino acid sequences of the three subunits of Ni-protein I were determined by Edman degradation and used to identify the encoding genes in a metagenome library of the mats. The codon usage and tetranucleotide signature of the three genes in the cluster mcrBGA revealed that the three genes are located on the genome of the dominant ANME-1 archaeon present in the mats. The deduced amino acid sequences show a high degree of sequence similarity to MCR from methanogenic archaea but with some distinct differences: in the α-subunit the glutamine, which in MCR from most methanogeninic archaea is post-translationally methylated at C2, is substituted by a valine (Figure 4). Two amino acids downstream of the valine there is a cysteine-rich sequence CCX4CX5C not present in MCR from methanogenic archaea (Figure 4). This cysteine-rich stretch in the α-subunit is also found in the DNA sequence of the gene homologues present in the metagenomic library of other microbial consortia catalyzing AOM [41]. Due

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Figure 4. Active site structure of methyl-coenzyme M reductase from Methanothermobacter marburgensis. The amino acids remarkably different in the enzyme (Ni-protein I) from methanotrophic archaea are highlighted.

to these differences and the presence of a modified F430 the catalytic properties of the enzyme from methanotrophic archaea could differ significantly from those of MCR from methanogenic archaea. Thus, the catalytic efficiency could be higher [37]. As indicated above, the most abundant MCR in microbial mats catalysing AOM in the Black Sea contains a modified F430 with a molecular mass of 951 Da [42] which has been identified to be 172-methylthio-F430 (S. Mayr and B. Jaun, 2006, unpublished). This 951 Da cofactor, which can easily be identified by its characteristic MALDI-TOF mass spectrum, was not found in any of the methanogenic archaea analyzed in this respect nor in microbial cells present in the anaerobic digesters of the waste treatment plant in Marburg. The modified cofactor was found, however, in all habitats with AOM. But, besides the modified cofactor, the normal F430 with a mass of 905 Da is always present. Met. Ions Life Sci. 2, 323–356 (2007)


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It has already been mentioned that Ni-protein II, isolated from the microbial mats catalyzing AOM in the Black Sea, contained only the 905 Da cofactor [42], indicating that AOM is not restricted to MCR containing the 951 Da cofactor [37].

4.

CATALYTIC PROPERTIES OF METHYL-COENZYME M REDUCTASE

Besides the reduction of methyl-coenzyme M with coenzyme B (Figure 2) methyl-coenzyme M reductase also catalyzes the reduction of ethyl-coenzyme M and of 3-(methylthio) propionate with coenzyme B and the reduction of methylcoenzyme with N-6-mercaptohexanoyl threonine phosphate albeit at less than 1% of the catalytic efficiency [87]. Reduction of ethyl-coenzyme M with coenzyme B has been shown to proceed with inversion of stereo configuration [132]. Reversible inhibitors are allyl-coenzyme M, propyl-coenzyme M, coenzyme M, N-6-mercaptohexanoylthreonine phosphate, N-8-mercaptooctanoylthreonine phosphate, N-nonanoylthreonine phosphate, N-6-(methylthio)hexanoylthreonine phosphate, and N-7-(methylthio)heptanoylthreonine phosphate. Suicide inhibitors are 2-bromoethanesulfonate, 3-bromopropionate, cyano-coenzyme M, seleno-coenzyme M, and trifluoromethyl-coenzyme M, which convert active MCRred1 into inactive EPR silent forms, and 3-bromopropanesulfonate, 3iodopropanesulfonate and 4-bromobutyrate, which convert active MCRred1 into an inactive form exhibiting the MCRBPS EPR signal [87,121]. Inactivation of MCRred1 by 2-bromoethanesulfonate, 3-bromopropionate, cyano-coenzyme M, seleno-coenzyme M, and trifluoromethyl-coenzyme M was found to be dependent on the presence of coenzyme B [87]. Substrate specificity and inhibitor specificity are consistent with the structure of the active site, as revealed from the crystal structure of inactive MCRox1-silent and MCRsilent (see Section 3.1). The mechanism of the suicide inhibition can be explained by the reactivity of Ni(I)F430 towards the electrophilic inhibitors (see Section 3.2).

4.1.

The Reversibility of the MCR-Catalyzed Reaction

The MCR-catalyzed reaction is exergonic (Figure 2). Under standard conditions (nongaseous substrates and products at 1 M concentration and CH4 at 105 Pa pressure) the free energy change (∆Go) associated with the reaction is estimated to be 30 kJ/mol methane (see below). The free energy change under physiological conditions (∆G) is obtained from ∆G ∆Go RTln [Products]/[Substrates] 30 5.7 log [Products]/[Substrates]. The equation predicts that the back reaction becomes exergonic when the product to substrate concentration ratio is approximately Met. Ions Life Sci. 2, 323–356 (2007)

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105. Such a ratio is physiologically not unrealistic. At 105 Pa methane the ratio is 105 when the intracellular concentration of CoM-S-S-CoB is, e.g., 1 mM (103 M) and that of CH3-S-CoM and HS-CoB each 0.1 mM (104 M). Consistent with this, methanogenic archaea have been shown to be capable of very slow methane oxidation [133,134]. In some of the earlier reports [135,136] it has to be considered, however, that the 14C methane used to follow AOM was generated from 14CO2 by methanogens and was therefore most likely contaminated by 14CO [137]. ∆Go for the MCR-catalyzed reaction is obtained from the difference in the free energy changes associated with several reactions [4,138]. One of these reactions is the reduction of CoM-S-S-CoB with H2. ∆Go for this reaction was revised recently due to the finding that the redox potential of the CoM-S-S-CoB/ HS-CoB HS-CoM couple is 143 ± 10 mV rather than 200 mV [139]. As a result, ∆Go for methyl-coenzyme M reduction with coenzyme B decreased from 45 kJ/mol to 33 kJ/mol. This value also has some uncertainty since it is in part based on ∆Go 28 kJ/mol associated with methyl-coenzyme M formation from methanol and coenzyme M, which was calculated from differences in bond energies which neglects differences in solvation energies [138]. ∆Go for methyl-coenzyme M reduction is probably best given as being 30 ± 10 kJ/mol.

4.2. The Catalytic Efficiency of Methyl-Coenzyme M Reductases Methyl-coenzyme M reductase (isoenzyme I) from Methanothermobacter marburgensis catalyzes methane formation from methyl-coenzyme M with a maximal specific activity of approximately 100 U ( µmol/min)/mg protein (500 s1) [4]. Km values for methyl-coenzyme M and coenzyme B are approximately 1 and 0.1 mM, respectively [11]. Exponentially growing M. marburgensis can produce methane at a specific rate of up to 5 U/mg protein. Consistently, such grown methanogenic archaea contain MCR at concentrations of 5–10% of the soluble cell proteins. Due to experimental reasons (equilibrium already reached after a few turnovers) the rate of the back reaction catalyzed by the enzyme has not yet been determined experimentally. The specific rate can be estimated, however, employing the Haldane equation, which correlates the equilibrium constant (Keq) of a reaction with the catalytic efficiency (kcat/KM) of an enzyme to catalyze the forward and the back reaction: Keq catalytic efficiency (forward reaction)/catalytic efficiency (backwards reaction). Keq for the MCR catalyzed reaction is calculated from ∆Go RTlnKeq 30 kJ/mol to be near 105. Assuming the KM values of MCR for its substrates and products to be all, e.g., 0.1 mM, the Haldane equation predicts that MCR with a Vmax for methyl-coenzyme M reduction of 100 U/ mg catalyzes methane oxidation at a maximal specific rate of 1 mU/mg (105 100 U/mg). As indicated above, ∆Go 30 kJ/mol of the MCR catalyzed reaction is only known with an uncertainty of ±10 kJ/mol. Therefore, the maximal specific rate of methane oxidation could be as high as 10 mU/mg and as low as 0.01 mU/mg. Met. Ions Life Sci. 2, 323–356 (2007)


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4.3. The Catalytic Mechanism of Methane Formation The active site structures of MCRox1-silent and MCRsilent reveal that methylcoenzyme M reduction to methane takes place in a hydrophobic pocket from which water is almost completely excluded. Therefore, when entering the active site via the hydrophobic channel methyl-coenzyme must be stripped of water and after the reaction the hydrophobic heterodisulfide has to be expelled into the water phase. Of the two substrates methyl-coenzyme M must enter the 50-Å-long channel first since when coenzyme B is bound the substrate channel is completely locked. Methyl-coenzyme M enters with its sulfonate group (probably protonated) ahead since once entered the molecule can no longer turn around. Methyl-coenzyme M can become positioned in the cavity above F430 such that either its methyl group or its thioether sulfur points towards the Ni(I) of F430 (Figure 5). Coenzyme B, which enters second, binds in a manner such that its thioheptanoyl group points towards F430 and the phosphate moiety towards the entrance of the channel. The sulfur of coenzyme B gets positioned above the nickel of F430 at a distance of 8 Å, which is too far for the thiol group to directly interact with the nickel. It can, however, come within van der Waals’ distance to the methyl group or the thioether sulfur of methyl-coenzyme M dependent on whether methyl-coenzyme M is in contact with the Ni(I) via its thioether sulfur or its methyl group carbon, respectively. The catalytic cycle most probably starts with a rate-limiting conformational change within the active site, which is induced upon binding of coenzyme B and which forces methyl-coenzyme M and Ni(I) of the prosthetic group to interact in the active enzyme MCRred1. This is deduced from the finding that inactivation of MCRred1 by bromoethanesulfonate and by other suicide inhibitors is dependent

Figure 5. Optimal position of methyl-coenzyme M in the active site of methyl-coenzyme M reductase assuming (A) catalytic mechanism 1 and (B) catalytic mechanism 2. The long aliphatic arm of coenzyme B can reach into the channel only to the extent where its terminal thiol group still is in a distance of 8 Å from the Ni(I). Met. Ions Life Sci. 2, 323–356 (2007)

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on the presence of coenzyme B [87] and that the substrate analog coenzyme M binds only with its thiol group to Ni(I) in the presence of coenzyme B as revealed by EPR spectroscopy [123,124]. In single turnover experiments methane formation from methyl-coenzyme M was found to be dependent on coenzyme B [140]. Steady-state kinetics have revealed a ternary complex catalytic mechanism [11]. Two different mechanisms for methyl-coenzyme M reduction have been proposed, both of which involve 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 [141,142]. The redox potential Eo of the thiyl radical/thiol couple has been estimated to be of the order of 1.3 V compared with NHE [142] and that of the disulfide/disulfide radical anion couple to be 1.4 V [143]. They are similar to the redox potentials of the Ni(III)F430 /Ni(II)F430 couple (Eo 1 V) and of the Ni(II)F430 /Ni(I)F430 couple (Eo 0.6 V) (see Sections 2.2, 2.3 and 2.4) In mechanism 1 the methyl group of methyl-coenzyme M reacts with the Ni(I) (Figure 5A) in a nucleophilic substitution reaction yielding methyl-Ni(III) and coenzyme M, which in turn react to methyl-Ni(II) and the thiyl radical of coenzyme M. Methyl-Ni(II) then reacts with a proton in an electrophilic substitution reaction to methane and Ni(II) and the coenzyme M thiyl radical reacts with coenzyme B yielding a disulfide anion radical, which is a strong reductant and which reduces Ni(II) back to Ni(I) thus closing the catalytic cycle [108,110,111]. This mechanism is mainly supported by the findings that MCR-catalyzed ethylcoenzyme M reduction proceeds with inversion of stereo configuration at C1 of its ethyl group [132], that enzyme bound Ni(I)F430 reacts with 3-bromopropane sulfonate to an alkyl-Ni(III) ↔ alkyl-radical Ni(II) species (see Section 3.4) and that free Ni(I)F430M in aprotic solvents reacts with methyl iodide to methyl-Ni(II)F430, which subsequently can be protonolyzed to methane and Ni(II)F430 [46]. 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 consistent with a nucleophilic substitution as first step in the catalytic cycle [121]. However, based on density functional calculations, Pelmenschikov et al. have concluded that mechanism 1 would be energetically unfavorable [144,145]. As an alternative they proposed mechanism 2, in which the thioether sulfur in methyl-coenzyme M is attacked by Ni(I) (Figure 5B) according to a radical substitution yielding a Ni(II) thiolate and a free methyl radical which in turn reacts with HS-CoB yielding methane and a CoB-Sᠨ thiyl radical. The latter reacts with coenzyme M thiolate to the disulfide anion radical which, like in mechanism 1, is used to re-reduce the Ni(II)F430 to Ni(I)F430. Mechanism 2 is backed up by the experimental finding that in active MCR coenzyme M reversibly coordinates with its thiol sulfur to Ni(I) of F430 when coenzyme B is present (see Section 3.3). This mechanism is not supported by the finding that methyl-coenzyme M reductase catalyzes the reduction of ethyl-coenzyme M with only very low catalytic efficiency and does not at all catalyze the reduction of allyl-coenzyme M although allyl-coenzyme M binds to the enzyme and competitively inhibits it with a Ki of Met. Ions Life Sci. 2, 323–356 (2007)


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0.1 mM [87]. Release of a free allyl radical according to mechanism 2 should be much more favorable than formation of a free methyl radical. Considering an involvement of methyl-coenzyme M reductase in the anaerobic oxidation of methane, both mechanisms are not very attractive. Assuming mechanism 1, methane oxidation would start by an insertion of Ni(II)F430 into a CH bond of methane with concomitant release of a proton. This is highly unlikely since Ni(II) of F430 is not electrophilic enough to be able to attack methane with a pKa of above 40. The low electrophilicity of Ni(II)F430 is reflected in the low redox potential Eo 0.6 V of the Ni(II)F430 /Ni(I)F430 couple [46,77]. (Only a reaction of methane either end-on or side-on with Ni(III)F430 as described for the activation of CH bonds by other high-valent metal complexes can be envisaged [146]). Mechanism 2 is likewise problematic. Methane oxidation would start by the reaction of methane with the CoB-S• thiyl radical. The bond dissociation energy of the CH bond in methane is 439 kJ/mol compared with that of the SH bond of only 365 kJ/mol, which makes a reaction of methane with a thiyl radical yielding a methyl radical and a thiol thermodynamically very unfavorable.

4.4.

Indications for a Dual-Stroke Engine Mechanism

There is evidence that the two active sites are structurally and functionally interlinked. The crystal structure (see Section 3.1) revealed that the two α subunits in the enzyme are intertwined such that a conformational change in the one active site (made up of the subunits α, α, β and γ) can be transmitted to the other active

Figure 6. Dual-stroke engine mechanism proposed for methyl-coenzyme M reductase. The mechanism allows the coupling of endergonic steps of the catalytic cycle in the one active site to exergonic steps in the other active site. The coupling is predicted to lower the activation energy both in the forward and the back reaction [121]. Met. Ions Life Sci. 2, 323–356 (2007)

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site (made up of the subunits α, α, β and γ) and vice versa. An indication for the coupling of the two active sites is the finding that at most 50% of the enzyme are converted from the MCRred1 state into the MCRred2 state upon addition of coenzyme M and coenzyme B (see Section 3.3). MCR shows ‘half-of-the-sites’ reactivity. Based on these findings it has been proposed that the enzyme operates according to a dual stroke engine mechanism [121] (Figure 6). This would allow the coupling of endergonic steps of the catalytic cycle in one active site to the exergonic steps in the other site. The mechanism predicts that MCRred1/silent with one active site in the red1 state and the other in an EPR silent state should be inactive. Indeed, some MCR preparations with relatively high-spin concentrations of the red1 signal show relatively low catalytic activity [121].

4.5. The Anaerobic Oxidation of Methane The presence of high concentrations of methyl-coenzyme M reductase in methanotrophic archaea strongly suggests that this enzyme catalyzes the first step in the anaerobic oxidation of methane. The thermodynamics of the MCR-catalyzed reaction and the catalytic properties of MCR from methanogenic archaea appear to conform to this proposed function. But, since the free energy change associated with the MCR-catalyzed reaction and the maximal specific rate of AOM are not yet known with certainty, it is too early for a decisive conclusion.

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

ABBREVIATIONS AOM Cdh CH3-S-CoM CoM-S-S-CoB DFT Eha Ehb ENDOR EXAFS

anaerobic oxidation of methane carbon monoxide dehydrogenase methyl-coenzyme M heterodisulfide of coenzyme M and coenzyme B density function theory energy-converting [Ni-Fe]-hydrogenase isoenzyme I energy-converting [Ni-Fe]-hydrogenase isoenzyme II electron nuclear double resonance extended X-ray absorption fine structure spectroscopy

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F F430M Frh Hdr Hmd HOMO HS-CoB HS-CoM HYSCORE LUMO MCD MCR Mcr MO Mrt Mtd Mvh NHE SCE XANES XAS

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Faraday constant pentamethylester of the nickel corphin coenzyme F430 F420-reducing [Ni-Fe]-hydrogenase heterodisulfide reductase iron–sulfur-cluster-free hydrogenase highest occupied molecular orbital coenzyme B coenzyme M hyperfine sublevel correlation lowest unoccupied molecular orbital magnetic circular dichroism methyl-coenzyme M reductase isoenzyme I from Methanothermobacter marburgensis methyl-coenzyme M reductase isoenzyme I molecular orbital methyl-coenzyme M reductase isoenzyme II F420-dependent methylenetetrahydromethanopterin dehydrogenase F420-nonreducing [Ni-Fe]-hydrogenase normal hydrogen electrode standard calomel electrode X-ray absorption near-edge structure X-ray absorption spectroscopy

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9 Acetyl-Coenzyme A Synthases and Nickel-Containing Carbon Monoxide Dehydrogenases Paul A. Lindahl1 and David E. Graham2 1

Departments of Chemistry and of Biochemistry and Biophysics, Texas A&M University, College Station, TX 77843, USA 2

Department of Chemistry and Biochemistry, The University of Texas at Austin, Austin, TX 78712, USA <[email protected]>

1. INTRODUCTION 2. STRUCTURE AND FUNCTION OF CARBON MONOXIDE DEHYDROGENASES 2.1. Redox Properties of the C-Cluster 2.2. Role of Sulfide Ion Bridging Ni and Feu 2.3. Nickel-Deficient Precursor C-Cluster 2.4. Nickel Incorporation in vivo 3. SEQUENCE ANALYSIS AND PHYLOGENY OF CARBON MONOXIDE DEHYDROGENASES 4. ACETYL-COENZYME A SYNTHASES/CARBON MONOXIDE DEHYDROGENASES 4.1. The Aox State 4.2. The Ared-CO State and the Heterogeneity Problem 4.3. Reductive Activation 4.4. Methylated and Acetylated States 4.5. Alternative Mechanisms 4.6. Connection to the β Subunits Metal Ions in Life Sciences, Volume 2 © 2007 John Wiley & Sons, Ltd


358 361 364 366 366 368 369 373 375 375 377 379 380 380

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5.

SEQUENCE ANALYSIS AND PHYLOGENY OF THE α SUBUNIT 6. CORRINOID IRON–SULFUR PROTEINS 7. ACETYL-COENZYME A DECARBONYLASE/SYNTHASES 8. PHYSIOLOGICAL ROLES AND EVOLUTION OF ACETYL-COENZYME A SYNTHASE/CARBON MONOXIDE DEHYDROGENASE PROTEINS 8.1. The Archaeal ACDSs 8.1.1. Methanobacteriales 8.1.2. Methanomicrobiales 8.1.3. Methanococcales 8.1.4. Methanopyrales 8.1.5. Methanosarcinales 8.1.6. Archaeoglobales 8.2. ACS/CODH Genes in Bacteria 8.2.1. Clostridia 8.2.2. Proteobacteria 9. ORIGINS AND EVOLUTION OF ACDS, ACS/CODH, AND CODH COMPLEXES ACKNOWLEDGMENTS ABBREVIATIONS APPENDICES Appendix 1. Spectroscopic properties of the C*-cluster Appendix 2. Alignment of β Subunit Protein Sequences Appendix 3. Alignment of α Subunit Protein Sequences Appendix 4. Alignment of γ Subunit Protein Sequences Appendix 5. Alignment of δ Subunit Protein Sequences Appendix 6. Protein Sequence Identifiers and Accession Numbers REFERENCES

1.

381 382 384

386 386 386 386 387 387 388 389 389 389 391 392 393 394 395 395 396 402 405 407 409 411

INTRODUCTION

This chapter focuses on a diverse group of oxygen-sensitive nickel-containing enzymes which have in common the ability to catalyze Reaction (1), the reversible oxidation of CO to CO2. Basic properties of these enzymes are given in Table 1. CO H 2 O CO2 2 e 2 H + Met. Ions Life Sci. 2, 357–416 (2007)

(1)

ACETYL-CoA SYNTHASES AND Ni-CONTAINING CODH

Table 1.

359

Subunit features of CODH, ACS/CODH, and ACDS proteins.

Name Physiological functions

E.C. Numbers Prototypes

Subunit αa Subunit βa

Subunit γb

Subunit δb Subunit ε Nickel-insertion accessory proteinc

CODH

ACS/CODH

ACDS

Carbon monoxide dehydrogenase

Acetyl-CoA synthase/ carbon monoxide dehydrogenase

Acetyl-CoA decarbonylase/ synthase

Carbon monoxide oxidation

Acetogenesis or acetate oxidation

Acetyl-CoA synthesis or acetate cleavage in methanogenesis 1.2.99.2 1.2.7.4 1.2.7.4 2.3.1.169 2.3.1.169 Rhodospirillum Moorella thermoacetica Methanosarcina rubrum thermophila (ACS/CODHMt) (CODHRr) Carboxydothermus hydrogenoformans (ACS/CODHCh) Not present 80 kDa 55 kDa Contains A-cluster Contains A-cluster 85 kDa 70 kDa 70 kDa Contains B- and Contains B- and Contains B- and C-clusters C-clusters C-clusters Shares D-cluster with Shares D-cluster with second β subunit second β subunit Contains [Fe4S4] Eand F- clusters Not present 50 kDa 50 kDa Contains [Fe4S4] cluster Contains [Fe4S4] cluster Not present 35 kDa 45 kDa Contains cobalamin Contains cobalamin Not present Not present 20 kDa 28 kDa 28 kDa 29 kDa

Subunits were assigned Greek letters based on their molecular masses, with that of α β γ δ ε. This nomenclature has created some confusion in the literature because the highest molecular mass subunit in the acetogenic enzyme is not the highest molecular mass subunit in the methanogenic enzyme. For this review, we use the acetogenic subunit nomenclature for all enzymes to facilitate comparisons. b In the ACS/CODH subunits γ and δ comprise an autonomous complex and are sometimes referred to as the α and β subunits of the CoFeSP complex. c The putative nickel insertion accessory protein CooC (or AcsF) is frequently associated with gene clusters encoding β subunits. This protein’s precise function is unknown. a

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One class of enzymes catalyzes only this reaction. These so-called carbon monoxide dehydrogenases (CODHs) are monofunctional β2 homodimers. Another class additionally catalyzes Reaction (2), the synthesis of acetyl-coenzyme A. CH 3 -Co3FeSP CO CoASH CH 3 -C(O)-CoA Co1FeSP H (2) These bifunctional α2β2 enzymes are called acetyl-CoA synthases/carbon monoxide dehydrogenases (ACS/CODH’s). For these enzymes, the methyl group is donated by an autonomous heterodimeric corrinoid–iron–sulfur protein (CoFeSP). The methyl group originates from CH3-tetrahydrofolate, and is transferred onto CoFeSP according to Reaction (3). CH 3 -H 4 F Co1FeSP H CH 3 -Co3FeSP H-H 4 F

(3)

The third class of enzymes also catalyzes the synthesis of acetyl-CoA, but the methyl group is transferred directly from CH3-tetrahydrosarcinapterin (H4SPT) or related molecules, as shown in Reaction (4). CH 3 -H 4 SPT CO CoASH CH3 -C(O)-SCoA H-H 4 SPT

(4)

The quaternary structure for these enzymes is (αβγδε) n (where n is even, possibly equal to 8) in which the γ and δ subunits are homologous to CoFeSP [1]. They generally operate reversibly and often function to decarbonylate acetyl-CoA according to Reaction (5) rather than to synthesize it; thus they are called acetylCoA decarbonylase/synthases (ACDS’s). CH 3 -C(O)-SCoA H 4 SPT CH3 -H 4 SPT COCoASH

(5)

The CO is then oxidized to CO2 and the methyl group of CH3-H4SPT is ultimately converted into CH4. In the past few years, a dozen review articles have focused on these enzymes! This clearly reflects the high level of interest generated most recently by the remarkable insights gained from a series of X-ray crystal structures reported during the period 2001–2004 [2–6]. These 4 years represent somewhat of a watershed; so much changed or was clarified during this period that the years prior now seem like the dark ages! Reviews by Drennan et al. [7] and Volbeda and Fontecilla-Camps [8–10] stress these structural advances. The arrival of these structures immediately stimulated activity in modeling the novel active sites of these enzymes, and reviews by Riordan [11], Hegg [12], Evans [13], and Harrop and Mascharak [14] highlight these advances. The structures also prompted advanced computational and spectroscopic investigations, and these are described by the review of Brunold [15]. Perhaps more so than for any other subdiscipline of this field, these new structures enriched our understanding of the enzymes’ Met. Ions Life Sci. 2, 357–416 (2007)


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mechanisms; these aspects are highlighted by Grahame [16], one of us [17] and by Ragsdale [18], who perhaps more than anyone has been responsible for ‘raising and nurturing’ these enzymes in those early years. Given this recent flurry of activity, we hesitate to write yet another review. We noticed that no review since 2001 [19] has emphasized the evolutionary relationships of these enzymes, even though the number of relevant genes sequenced is far greater now than it was in 2001 due to microbial genome sequencing projects. As such, in this review we decided to combine these aspects with the current understanding of the structure, spectroscopic properties, and mechanistic enzymology. We assume some general background knowledge of these enzymes, much of which can be found in previous reviews, and emphasize recent developments that have not yet been summarized in a review. We also highlight the most controversial aspects regarding these enzymes that continue to evade resolution.

2. STRUCTURE AND FUNCTION OF CARBON MONOXIDE DEHYDROGENASES Virtually all of our knowledge of these enzymes has been obtained by studying the CODHs from Rhodospirillum rubrum, Carboxydothermus hydrogenoformans, and the homologous β2 dimer of the ACS/CODH from Moorella thermoacetica. All are homodimeric β2 dimers with subunit masses between 60–70 kDa. X-ray crystal structures of all three enzymes (CODHRr, CODHCh, and CODHMt) have been determined [2–5] and, except for a few important differences (described below), their structures are equivalent. We will call enzymes with these structural features standard CODH’s. Each β2 dimer contains five metal centers of three different types (Figure 1, top structure). Each subunit contains 1 B- and 1 C-cluster while a single D-cluster bridges the two subunits. They are arranged in the shape of a “V” of the order C(βL) – B(βR) – D – B(βL) – C(βR) where βL and βR refer to different subunits, the D-cluster forms the base of the V, and each cluster is ⬃11 Å from the next. The C- and B-clusters are buried in the protein while the D-cluster is surface-exposed (Figure 1, top right structure). The B- and D-clusters are both [Fe4S4] 2 clusters. The B-cluster is a traditional [Fe4S4] 2/1 cluster, coordinated by 4 cysteinyl residues (see Table 2 for a list of conserved residues) and with a redox potential of E0 ⬃ 0.4 V compared with NHE. In the reduced state this cluster has an {S 1/2:S 3/2} spin state mixture where the majority S 1/2 portion exhibits an EPR signal with gave 1.94. The redox and magnetic properties of the D-cluster are uncertain. The cluster might remain in the 2 core oxidation state [20] or it might have redox properties similar to that of the B-cluster [21]. In any event, there is no doubt that the B- and D-cluster serve as an ‘electron wire’ to transfer electrons between external redox agents and the active site C-cluster, as required by Reaction (1). In Met. Ions Life Sci. 2, 357–416 (2007)

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Figure 1. Structure of CODHRr [3] (top) and ACS/CODHMt [5] (bottom). The structure of CODHRr on the right was generated by rotating that on the left by 90 around the axis running vertically through the center of the protein. The direction of rotation is such that the D-cluster extends towards the viewer in the structure on the left. These images as well as those of Figure 6 were generated using PyMOL. Table 2. Selected conserved residues of the β subunit. Residue Cys Cys Cys Cys Cys Cys His His His His His Cys Cys Cys Cys Cys Cys Lys

CODHMt

CODHRr

CODHCh

059 067 068 071 076 090 113 116 119 122 283 316 317 355 470 500 550 587

041 049 050 052 058 072 095 098 101 108 265 299 300 338 451 481 531 568

039 047 048 051 056 070 093 096 099 102 261 294 295 333 446 476 526 563

Met. Ions Life Sci. 2, 357–416 (2007)

Description Ligand to D-cluster Ligand to D-cluster Ligand to B-cluster Ligand to B-cluster Ligand to B-cluster Ligand to B-cluster Proton Transfer Network Proton Transfer Network Proton Transfer Network Proton Transfer Network Ligand to Feu of C-cluster Persulfide bond to [Fe3S4] subsite Ligand to Feu of C-cluster Ligand to [Fe3S4] subsite C-cluster Ligand to [Fe3S4] subsite C-cluster Ligand to [Fe3S4] subsite C-cluster Ligand to Ni of C-cluster Proton transfer network


363

standard CODHs, the ligands coordinating the D-cluster are cysteines, two from each subunit. Reaction (1) also involves the transfer of protons into/out of the C-cluster. As first suggested by Doukov et al. [4] and later confirmed by the site-directed mutagenesis study of Kim et al. [22] there is a network of conserved residues that allows protons to migrate between the C-cluster and external proton acceptors/ donors in solution. For most enzymes, the residues involved include 4 His and perhaps Lys and an Asn (see Table 2). The pathway used for proton transfer is distinct from that used for electron transfer. There are different patterns of these basic amino acids (see Table 3 in Section 3 and Appendix 2). The C-cluster consists of the structure shown in Figure 2. It can be viewed as an [Fe3S4] subsite linked to a [Ni…Fe] subsite. The Fe3S4 subsite is coordinated by three cysteinate sulfurs and has three µ3-bridging facial sulfides that coordinate the [Ni…Fe] subsite. The unique Fe, hereafter referred to as Feu, is coordinated by one of these sulfides and by a cysteinyl sulfur and histidinyl nitrogen. The Ni is coordinated to the remaining two µ3-bridging facial sulfide ions as well as to a cysteinyl sulfur to afford a T-shaped trigonal planar geometry. There is some variation in the exact location of the Ni and Feu, suggesting conformational heterogeneity and/or the possibility that minor structural changes between these two ions occur during catalysis [5]. There are two open binding sites, one exclusively on the Ni and the other that bridges to Feu (called E1 and E2 [10]). These are the uncontroversial portions of the C-cluster structure, in that all reported structures possess them. Other features are present in some structures and absent in others, raising questions as to which form represents functional C-clusters. Before we discuss these aspects, however, we need to consider the redox properties of the C-cluster.

Figure 2.

Structural model of the C-cluster. Residue numbers refer to CODHMt. Met. Ions Life Sci. 2, 357–416 (2007)

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Figure 3. Redox properties of the C-cluster.

2.1.

Redox Properties of the C-Cluster

The C-cluster can be stabilized in four oxidation states, called Cox, Cred1, Cint, and Cred2 (Figure 3). Cox is diamagnetic and with no Feu evident by Mössbauer spectroscopy. The diamagnetism and Mössbauer parameters (isomer shift δ and quadruple splitting ∆ EQ) suggest formal valences of {2Fe2 and 1Fe3} for the irons of the [Fe3S4]1 subsite and Fe3 for (unresolved) Feu [21,23]. Reduction to the S 1/2 Cred1 state occurs with a redox potential of E0 ⬃ 0.1 V. In this state, Feu is high-spin 2 according to Mössbauer spectra of Cred1, suggesting that the added electron goes to Feu. ENDOR spectra of this state reveal a strongly coupled proton, suggesting hydroxyl coordination to the cluster [24]. Based on these results and those of Hu et al. [23], reduction to the Cred1 state appears to follow Reaction (6) where the hydroxyl group bridges Feu2 and Ni2 [21]. Cox {[ Fe3S4 ]1[ Ni 2 … Fe3 ]}1e H 2 O Cred1{[ Fe3S4 ]1[ Ni 2 -O(H)-Fe 2 ]}H

(6)

H2O may already be bound in the Cox state, but there is no evidence for (or against) this. Ti(III) citrate causes reduction to the Cred1 state into an EPR-silent redox state called Cint [25]. This state appears to be one electron more reduced than Cred1. Using CO or other low-potential reductants (such as dithionite), the Cred1 state can be reduced by 2 electrons to another S 1/2 state called Cred2 in Reaction (7). Cred1{[ Fe3S4 ]1[ Ni 2 -O(H)-Fe 2 ]}2 e H Cred2 {[ Fe3S4 ]1[“ Ni 0 ”… Fe 2 ]}H 2 O Met. Ions Life Sci. 2, 357–416 (2007)

(7)


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ENDOR spectra suggest that the hydroxyl group is absent in the Cred2 state [24]. The assignment of “Ni0” should not be taken literally as we use it here only as an electronic bookkeeping device. Indeed the location of the added electrons is uncertain. Recently, Gu et al. [26] have performed Ni K-edge XAS on a CODH from Carboxydothermus hydrogenoformans in what appears to have been the Cox and Cred2 states. In the Cox state, the Ni appeared to be in the 2 state, with square planar geometry (4 S ligands at 2.20 Å). In the Cred2 state, the Ni remains 2, but there is a change in geometry, lengthening of the Ni–S bond lengths (to 2.25 Å), and evidence of a Ni–Fe interaction at 2.7 Å. These results suggest a structural change occurring upon conversion to the Cred2 state, but no change in Ni oxidation state. Volbeda and Fontecilla-Camps [9] proposed that the two electrons might be stored as a hydride coordinated between the Ni and Feu. The only concern here is the absence of a strong proton ENDOR signal in the Cred2 state, which would seem likely if such a species existed. One might also propose that the two electrons localize on the [Fe3S4] subsite, but this subsite appears to be in the 0 state (2 Fe2 and 1 Fe3) when the entire cluster is in the Cred1 state, and an additional 2 electrons would reduce it to a state that seems unachievable (2 Fe2 and 1 Fe1). One electron could localize on the [Fe3S4] subsite and the other on the Ni, but in this case, it seems that the magnetic properties of the resulting {[Fe3S4] 0 [Ni1…Fe2]} state would be substantially different from that of Cred1 [21]. In fact, Cred1 and Cred2 EPR signals are very similar, suggesting similar electronic states. A final possibility involves a persulfide bond that was observed in one (and only one) crystal structure of the enzyme between Cys316 of ACS/CODHMt (Appendix 2) and the µ3-bridging sulfides that coordinate Feu in CODHMt [5]. Supporting the importance of Cys316 is that it is largely conserved (Table 2 and Appendix 2). The idea would be that this bond is reduced in Cred2 and oxidized in Cred1. However, the C-cluster appears to be absent in a mutant enzyme lacking this cysteine, suggesting that this persulfide plays a role in either assembling or stabilizing the cluster [22]. During catalysis, CO reacts with the C-cluster in the Cred1 state while CO2 reacts with that in the Cred2 state, as shown in Reaction (8) (focusing only on the [Ni…Fe] subsite). [ Ni 2 -O(H)-Fe 2 ]CO [ Ni 2 -CO HO-Fe 2 ] [ Ni 2 -COOH Fe 2 ] [“ Ni 0 ” Fe 2+ ]CO2 H

(8)

During CO oxidation catalysis, electrons are transferred one at a time to the B-cluster (and then to the D-cluster and then to the protein exterior), generating the Cint state after the first electron transfer and Cred1 after the second, thereby completing the catalytic cycle. The step at which the substrate H2O bind to the C-cluster is unknown. Met. Ions Life Sci. 2, 357–416 (2007)

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2.2.

LINDAHL and GRAHAM

Role of Sulfide Ion Bridging Ni and Feu

Some structures, under some redox-states, additionally contain a µ2-bridging sulfide ion or a bridging cysteinate coordinating the Ni and Feu. The function of this sulfide ion is controversial. Dobbek et al. [2] have reported that it is critical for catalysis such that only the CODHCh structure, which contained this species, represents the active enzyme. The only obvious difference in the crystallization conditions between the three proteins was the presence of CO in both CODHRr and in ACS/CODHMt, and the absence of this substrate in CODHCh. When crystallized in CO, CODHCh lacked the sulfide bridge. Moreover, the activity of CODHCh was abolished after being incubated in CO for ⬃100 h [27]. They suggest that CO reacts with the sulfide to form COS and the inactive sulfide-free form of the enzyme. Using CODHRr, Feng and Lindahl [28] repeated the experiment of Dobbek et al. in which CODH activity was monitored in the presence and absence of CO. After an initial drop in activity, there was no loss of activity over the next 650 h! They also found that added sulfide ions partially inhibited catalysis. Sulfide bound exclusively to the Cred1 state but the enzyme could be reactivated by reduction to the Cred2 state. They suggested that HS displaces a bridging HO via Reaction (9) and that activation occurred via Reaction (10). {[ Fe3S4 ]1[ Ni 2 -O(H)-Fe 2u ]}HS {[ Fe3S4 ]1[ Ni 2 -S(H)-Fe 2 u ]} HO

(9)

{[ Fe3S4 ]1[ Ni 2 -S(H)-Fe 2u ]}2 e {[ Fe3S4 ]1[“ Ni 0 ”… Fe 2u ]}HS

(10)

Other anion inhibitors, including cyanide, azide, and thiocyanate behave similarly to sulfide, and are also proposed to bridge the Ni and Feu. Thus, controversy remains as to whether the sulfide ion bridging the Ni and Feu of the C-cluster is a dissociable inhibitor or a required component of active Cclusters.

2.3. Nickel-Deficient Precursor C-Cluster When R. rubrum is grown in the dark, under a CO atmosphere, and in Ni-deficient media, it synthesizes an inactive form of CODHRr (called CODHRr*) [29]. CODHRr* can be activated in vitro simply by adding millimolar concentrations of NiCl2 under reducing (approximately 0.4 V compared with NHE) conditions [30,31]. Mature C-clusters can also be assembled in E. coli [32], an organism Met. Ions Life Sci. 2, 357–416 (2007)


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that does not naturally contain a CODH and does not have chaperones specific for assembling C-clusters. However, heterologous expression of active CODH requires high levels of NiCl2 (0.5 mM) in the growth medium, and E. coli expresses a nickel active-transport system under anaerobic, Ni2-limiting conditions [33]. CODHRr* contains mature B- and D-clusters, but the C-cluster is present in a precursor form called C*. The structure of C* is controversial; it has been proposed to be both a [Fe4S4] 2/1 cluster [23] and a ‘preformed’ [Fe3S4]cluster spin-coupled to Feu, equivalent to the structure of the mature C-cluster without the Ni [20]. Recently, Jeon et al. [34] mutated the three cysteines that coordinate the [Fe3S4] subsite and used the precursor forms of these mutants to explore the mechanism of C-cluster assembly. Since C* is converted rapidly into the mature C-cluster by adding nothing more than Ni2 ions under reducing conditions, it would seem that the reaction is simply {C* Ni → C}, implying that C* consists of a structure with four Fe and four (or five) sulfide ions. We cannot exclude the possibility that the assembly reaction requires additional Fe or sulfide ions, and that, if these ions are not included in the reaction mixture, they might be scavenged from existing clusters within a population of protein molecules. Putative nickel chaperone proteins (discussed below) have been proposed to catalyze Ni insertion in vivo, although their reaction mechanisms are unknown. Jeon et al. [34] found that the first step in assembly is the reversible binding of Ni to reduced CODHRr* followed by an irreversible step which yields mature C-cluster. Two acid/base groups with pKa 7.6 and 9.5 are involved. The identity of the latter is uncertain, but the former may be the Cys531 cysteinate (which bridges the Ni and Feu in CODHRr and may bind only the Ni in CODHMt). His265 is also required for Ni binding (which is counterintuitive because it coordinates Feu, not the Ni). Assuming a preformed {[Fe3S4][…Feu]} model of C*, Jeon et al. [34] have proposed Mechanism A shown in Figure 4 (a second similar mechanism was also proposed in which a bridging sulfide ion is incorporated). We are puzzled by what appears to be a growing consensus favoring the preformed {[Fe3S4][…Feu]} model of C*, in that the evidence favoring the [Fe4S4] 2/1 model of C* cannot be easily ignored (Appendix 1). As such, we suggest an alternative mechanism of C-cluster assembly based on the [Fe4S4] 2/1 model (Figure 4, Mechanism B). Here, the Fe4S4 C*-cluster (along with the B-cluster and D-cluster) is assembled by the ubiquitous Fe/S-assembly machinery [35]. Two non-coordinating Cys thiols and a His residue are located along one Fe–S edge of the cubane, perhaps with the Fe of that edge uncoordinated (this could explain why some [Fe3S4] decomposition product was observed in CODHRr* under oxidizing conditions [23]. Aqueous Ni2 coordinates reversibly to 2 cubane sulfides. Upon reduction to the 1 core oxidation state, two Fe-µ4-sulfide coordinate bonds break, allowing that Fe to coordinate to His265 and Cys300. This promotes a conformational change in which the Fe and associated ligands rotate downward. One water from the aquated Ni ion coordinates Feu, forming a hydroxide Met. Ions Life Sci. 2, 357–416 (2007)

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Figure 4. Comparative mechanisms of C-cluster assembly. Residue numbers refer to CODHRr.

bridge. Only further experiments can differentiate the two mechanisms, but one advantage of Mechanism B is that it begins with a common metal center whose mechanism of assembly has been well studied, whereas Mechanism A begins with a unique metal center whose assembly is not explained.

2.4.

Nickel Incorporation in vivo

Although Ni2 can be incorporated into the C-cluster spontaneously in vitro, nickel insertion in vivo may require accessory proteins. During growth on CO, CODHRr binds a majority of the cell’s Ni pool [36], and these cells grow on medium supplemented with as little as 0.5 µM NiCl. The CODH gene cluster in this bacterium encodes CooC, CooT, and CooJ proteins that are proposed to catalyze Ni insertion in CODHRr. The function of CooT is unknown and a nonpolar cooT null mutation has little effect on CO-dependent growth [37]. The only identified homolog of cooT is in the related bacterium Rhodopseudomonas palustris str. BisB18. Null mutations in cooJ impair CO-dependent growth at low NiCl2 concentrations [37], although this putative nickel-binding protein also has homologs only in R. palustris BisB18. Met. Ions Life Sci. 2, 357–416 (2007)


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Most genomes that encode CODH, ACS/CODH or ACDS also encode homologs of the CooC nucleotide-binding protein, frequently in a gene cluster with other subunits. CooC and the homologous protein from M. thermoacetica (AcsF) are members of an enzyme family that includes nitrogenase iron protein (NifH) involved in FeMo cofactor formation, the septum site-determining protein (MinD), and the HypB and UreG proteins, which facilitate nickel insertion in hydrogenase and urease. These proteins share a conserved P-loop sequence at their amino-termini. Both CooC and AcsF exhibit low, but significant, levels of ATP phosphohydrolase activity [38,39]. The mechanism by which CooC inserts Ni is unclear. R. rubrum cooC mutants require ⬃650 µM NiCl2 in their growth medium for CO-dependent growth [37]. Yet the co-expression of AcsF in E. coli with heterologously expressed CODHMt did not enhance CODH activity; high levels of NiCl2 (500 µM ) were still required for CODH activation [39]. Neither purified CooC nor AcsF contain significant amounts of Ni2 or Fe2. Jeon et al. [34] observed that the addition of Ni2 and ATP to cell-free extract of R. rubrum grown in a low-nickel medium activated CODH activity. This effect was not seen in cooC null mutants or the K13Q variant of CooC that disrupts nucleotide binding. Yet no protein-specific activation of purified CODHRr* was observed using purified CooC, CooJ, ATP, and Ni2 [38]. Although the mechanism of CooC stimulation of CODH activity remains to be determined, a number of factors implicate CooC/AcsF in C-cluster assembly, including genetic studies in R. rubrum, evolutionarily conserved gene linkage to CODH in diverse organisms, the protein’s membrane association in R. rubrum and stimulatory activities in crude cell-free extracts.

3.

SEQUENCE ANALYSIS AND PHYLOGENY OF CARBON MONOXIDE DEHYDROGENASES

An alignment of forty β subunit protein sequences from diverse bacteria and archaea was prepared using the T-Coffee program (version 3.27) [40] with slight manual adjustment (Appendix 2). The alignment shows the conservation of metal cluster ligands, separated by regions of variable sequence and small insertions. The cysteine thiol that coordinates the C-cluster Ni is universally conserved, as is the histidine coordinating Feu. The three cysteine ligands of the [Fe3S4] portion of the C-cluster are almost universally conserved. The cysteine ligand to Feu (Cys317 in CODHMt) is conserved in most sequences, but is replaced by a glutamate residue in a cluster of four sequences, including A. fulgidus, M. acetivorans, D. hafniense, and C. hydrogenoformans (Table 3). A histidine residue replaces this cysteine in another paralog from Methanopyrus kandleri. We are uncertain how these changes affect the properties of the enzyme so we will refer to these as CODH-like sequences. Interestingly, Cys317Glu and Ser585Glu amino acid substitutions in several CODH-like proteins make their C-clusters Met. Ions Life Sci. 2, 357–416 (2007)

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Table 3.

LINDAHL and GRAHAM

Characteristics of CODHs and CODH-like proteins.

Organism

No. C-cluster

M. kandleri

1 2 3

A. fulgidus

2 3 1

B-cluster

D-cluster

Proton transfer pattern

Standard Feu (C→H); Per(C→T) Per(C→T)

Standard Absent

Standard Absent

(04)HXXHRHXXXNK (13)XXXXXXXXXXH

Standard

Standard

(06)YXHXENXXXQQ

Standard Feu (C→E); Per(C→I) Standard

Standard Standard

Standard C→S

(09)HXXHXHHXHHK (07)YXYHXXXTXEK

C→N

C→Q

(02)HXXHRHDHENK

M. acetivorans

2 5 3 4

Standard Standard Standard Feu (C→E); Per(C→I)

Standard Standard Standard Standard

Standard Standard Standard C→R

(02) (02) (01)HXXHXXXXXXK (07)

D. hafniense

1 2 3 4

Standard Standard Standard Feu (C→E); Per(C→I)

Standard Standard Standard Standard

Standard Standard Standard C→D

(03)HXXHXHHXXNK (04) (01) (12)YXHHXXXXXQK

C. hydrogenoformans

1 2 3 4 5

Standard Standard Standard Standard Feu (C→E); Per(C→I)

Standard Standard Standard Standard Standard

Standard Standard Standard Standard C→S

(03) (04) (01) (03) (07)

M. jannaschii

1 2

Standard Per(C→T)

Standard Standard

Standard C→Y

(05)HXGHRHHHXNK (06)

C. difficile

2 1

Standard Fe/S(C→D)

Standard Standard

Standard Standard

(08)HSDHRDHXXXK (06)

S. fumaroxidans

2 1

Standard Standard

Standard Standard

Standard C→S

(01) (01)

‘‘Feu’’, ‘‘Per’’ and ‘‘Fe/S’’ refer to the unique Fe, the cysteine involved in the persulfide bond, and the [Fe3S4] subsite, respectively. The nomenclature A→B indicates that in the designated cluster, ligand A has been replaced by ligand B. No. refers to the numbers indicated in Appendix 2. Proton transfer patterns (01–13) are designated by strings of conserved residues 113, 114, 115, 116, 118, 119, 121, 122, 125, 284, and 587 (M. thermoacetica numbers) with X indicating no conserved residue at that position.

more similar to the modified [Fe4S4] cluster of the Desulfovibrio desulfuricans hybrid cluster protein [41]. Because the function of the hybrid cluster protein is unclear, this association does little to help predict the functional implications of these substitutions. Met. Ions Life Sci. 2, 357–416 (2007)


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Other modifications seem to arise in concert, including at the D-cluster and at the cysteine (C316) involved in persulfide formation in CODHMt. Besides the four above-mentioned CODH-like sequences, this cysteine has also been mutated in three more distantly related sequences from Methanocaldococcus jannaschii, Clostridium difficile, and Methanopyrus kandleri. Each of these seven proteins has a functional β subunit homolog in their respective genomes, and these proteins have no known function. The non-conservative replacement of this cysteine residue by threonine or isoleucine suggests a change in catalytic function (neofunctionalization). The [Fe4S4] B-cluster comprises four cysteine residues, which are conserved in all sequences but two: homologs from Archaeoglobus fulgidus and Methanopyrus kandleri. The M. kandleri sequence has a deletion spanning all of the Bcluster cysteines, as well as the D-cluster cysteines. The [Fe 4S4] D-cluster, which is formed by two cysteine residues from each β subunit, is missing a cysteine in six of the seven sequences in the cluster of CODH-like genes. While it is conceivable that four of the amino acid replacements (Arg, Asp, Thr, and Tyr) could act as ligands, the presence of tryptophan in this position in a M. jannaschii homolog discounts that possibility. The [Fe4S4] E- and F-clusters are uniquely found in an inserted region of the archaeal ACDS sequences. The eight requisite cysteine residues are conserved in all of the ACDS sequences, except for the diverged M. kandleri homolog that is also missing the B- and D-cluster cysteines, described above. A series of basic residues is required to form a proton transfer network in the protein. While specific amino acids are poorly conserved in this feature, a periodic series of basic residues (especially histidine and arginine) could serve this purpose (Appendix 2 and Table 3). Because of the clear division between β subunits from ACDS and ACS/CODH or CODH enzymes, the sequence alignment was split into two subalignments that were used to infer the phylogeny of this gene. The phylogeny of the ACDStype proteins (Figure 5A) is mostly congruent with the small subunit ribosomal RNA phylogeny of these archaea [42] and the currently accepted taxonomy of the organisms [43]. This result suggests these were inherited vertically within the euryarchaea from a common ancestor that had a functional ACDS. An exception to this congruency is the clustering of the Methanothermobacter thermautotrophicus sequence with the Methanococcus maripaludis sequence, to the exclusion of the M. jannaschii sequence. Similar topologies in alternative trees and in the phylogenies of other subunits may indicate a horizontal gene transfer event, or a systematic artifact in the alignments or methods of phylogenetic inference. In several lineages, such as the Methanosarcinales, gene duplications have created paralogous β subunit genes. It is not known whether these paralogs have retained their ancestral function, lost function (nonfunctionalization), evolved

Met. Ions Life Sci. 2, 357–416 (2007)

ANME DNA

0.67

1.00 1.00

1.00

1.00 1.00

1.00 0.97 0.77

ANME DNA 1.00 0.1

A. ACDS β subunit phylogeny 1.00 1.00 1.00

1.00 1.00 1.00 1.00 1.00 1.00 1.00 0.76 0.99 0.98

1.00

1.00 1.00 1.00 1.00

1.00 1.00

1.00

1.00

0.1

B. CODH and ACS/CODH β subunit phylogeny

Figure 5. Phylogenies of the β subunit from ACDS complexes (A) and ACS/CODH or standard CODH proteins (B). The scale bar indicates 0.1 amino acid changes expected per site. Numbers near each interior branch are clade credibility values, the fraction of trees that contain each cluster of sequences shown in the consensus tree. Phylogenies were inferred using a Bayesian inference method implemented by the MrBayes program (version 3.1.2) [111], with four chains, the default mixed model of amino acid replacements with fixed rate matrices and a gamma distribution of rates approximated with four categories. Alternative phylogenies were inferred using the proml program or the protdist and neighbor programs (with 100 bootstrap replicates) [112]. Both programs used the Jones–Taylor–Thornton model of amino acid changes and assumed a gamma distribution of rates (α 2.4), approximated by three states. These trees are generally congruent with the trees shown here, and branches with high clade credibility values also have high bootstrap values.


373

to perform some of the ancestral gene’s functions (subfunctionalization), or evolved new functions (neofunctionalization). Answering these evolutionary questions will require additional studies of gene regulation and protein function. Genes encoding bacterial-type ACS/CODH β subunits or CODH have more complicated evolutionary histories (Figure 5B). Although the tree cannot be reliably rooted using the ACDS subunits, the phylogeny suggests that the ACS/CODH β subunit is not monophyletic with respect to standard CODH proteins. Either the standard CODH protein must have evolved from ACDS complexes in several steps, through the loss of associated subunits, or ACS/ CODH and ACDS complexes evolved through the parallel recruitment of other subunits. The latter model includes the possibility of non-orthologous gene displacement resulting from the horizontal transfer of β subunit genes. The modular nature of the ACS/CODH complex may facilitate these transfer events. ACS/CODH proteins are assigned functions based on biochemical characterization and gene cluster association. Some standard CODH proteins can be annotated based on their genes’ proximities to hydrogenase or other oxidoreductase genes. Another group of CODH genes offer no chromosomal gene context to identify a function. A phylogenetic cluster of seven genes including C. hydrogenoformans CODH-V has important differences in metal ligand residues (described above) that could alter its function. Two members of this CODHlike group, from M. jannaschii and M. kandleri, are chromosomally associated with three genes paralogous to the F420 -reducing hydrogenase subunits. The C. difficile homolog is also associated with iron–sulfur proteins of unknown function. On this basis, some CODH-like proteins might catalyze electron transfer reactions, but elucidating the nature of those reactions will require biochemical characterization.

4.

ACETYL-COENZYME A SYNTHASES/CARBON MONOXIDE DEHYDROGENASES

The best studied ACS/CODH is that from Moorella thermoacetica (previously Clostridium thermoaceticum). This 310 kDa bifunctional α2β2 tetramer catalyzes Reactions (1) and (2). Catalytic functions are largely independent, with β subunits responsible for CODH activity and α subunits responsible for ACS activity. In fact, recombinant α subunits can be expressed in E. coli such that after incubation in NiCl2, they exhibit ACS activity as long as CO (rather than CO2 /2e /2H) is used as a substrate [32,44,45]. For this reason, the α subunit is referred to as ‘ACS’, the β2 dimer as ‘CODH’, and the α2β2 tetramer as ‘ACS/ CODH’. Met. Ions Life Sci. 2, 357–416 (2007)

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Figure 6.

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Structural model of the A-cluster. See text for details.

The 82 kDa α subunit contains a single metal center called the A-cluster, the active site where acetyl-CoA is synthesized. Three crystal structures showing the A-cluster have been reported [4–6]; the ACS/CODHMt structure [5] is shown in Figure 1, bottom structure. The structure of active A-clusters (Figure 6) consists of an Fe4S4 cubane bridged (through a cysteinate) to a Ni ion (called the proximal Ni or Nip) which itself is bridged (through two cysteinates) to a second Ni ion (called the distal Ni or Nid). This form of the A-cluster has been observed in the open conformation of α , where Nip is located on the protein surface. Three protein domains can be recognized, with the one composed of the 30 kDa amino-terminus in contact with the β subunit. The structure of inactive A-clusters with Cu or Zn in place of Nip is also known [4]. Here, the geometry of the proximal site is tetrahedral, as is typical of complexes of the d10 ions Zn2 and Cu1. In the presence of inactive A-clusters, the conformation of α (called closed) is such that Nip and other portions of the A-cluster are located below the surface (Figure 7). The geometry of Nip in the open conformation is approximately square planar with three endogenous bridging cysteinyl ligands and an unidentified exogenous ligand. Nid has an S2N2 square planar geometry in both open and closed conformations. Besides the 2 bridging cysteinyl sulfurs, Nid is coordinated to two (probably deprotonated) amide nitrogens from the protein backbone. Based on mononuclear model complexes (discussed in [5]), this geometry and ligand environment favors a low-spin Ni2 valence, which makes it unlikely that Nid can be reduced beyond the 2 state.

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Figure 7. Spatial location of conserved residues in the α subunits of ACS/CODHMt. Residues conserved in all (or all but one) α subunit sequences, except for the six cysteines and one glycine known to coordinate the A-cluster, are plotted in color. Other residues are plotted in grey. The two subunits have been oriented independently to ease their comparison. Highlighted residues include E329, G330, E331, R334, E341, G344, E351, L352, G370, E398, E400, E404, R405, H408, Y413, G416, H419, Q422, R423, W427, R429, R488, D489, D497, E498, D501, Y504, Q510, S511, F512, A513, P514, V517, P522, D523, R524, G529, D535, G546, P547, K553, G562, N568, S575, P591, T593, S594, F598, E599, P606, E607, G610, G625, A632, G633, G637, G638, G643, F644, G646, K656, F657, D661, G662, G663, R666, V668, W669, P671, K675, I699, E702, F714, L715, H720, and P721.

4.1.

The Aox State

In the absence of a low-potential reductant like dithionite or Ti(III)citrate, the A-cluster is diamagnetic (S 0) with the electronic configuration Aox{[Fe4S4] 2Nip2Nid2} [17,46,47]. This so-called Aox state is incapable of accepting a methyl group from CH3-Co3FeSP or of participating in catalysis; it is an unactivated state [48,49].

4.2.

The Ared-CO State and the Heterogeneity Problem

When the enzyme is reduced and bound by CO, the A-cluster undergoes Reaction (11): A ox {[ Fe 4 S4 ]2 Ni 2p … Ni 2d}1e CO A red -CO{[ Fe 4 S4 ]2 Ni1pCO Ni 2d}

(11)

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Ared-CO is a S 1/2 state that exhibits the well-studied NiFeC EPR signal [18]. This signal has substoichiometric intensity, as it quantifies to only 0.1–0.3 spin/α [50]. Nip can be specifically removed by the bidentate chelator 1,10-phenanthroline [51]. A-clusters devoid of Nip are inactive and lack the NiFeC signal. However, Ni can be reinserted and activity restored by incubation in NiCl2. This lability is undoubtedly related to the ability of the proximal site to house different metal ions such as Cu and Zn. An early surprise was that only ⬃0.3 Ni/α can be removed or reinserted [52]. In conjunction with the low-spin concentrations of the NiFeC EPR signal, this result suggested that only ⬃30% of the A-clusters within a population of α subunits are catalytically functional while the remaining clusters are nonfunctional. This heterogeneity was subsequently confirmed by quantitative redox titrations [25] and Mössbauer spectroscopy [46]. Mössbauer spectra of a population of A-clusters poised in the Ared-CO state contain two species due to this heterogeneity. One represents ⬃30% of the A-clusters (the functional portion); it has S 1/2 and {[Fe4S4] 2Nip2-CO}. The other species represents ⬃70% of the A-clusters (the nonfunctional portion); it has S 0 and [Fe4S4] 2 clusters. The origin of the heterogeneity remains uncertain. It is not due to damage incurred during protein purification [53] nor does it depend on the lab in which the enzyme is isolated. Gauged from the low spin concentration of the NiFeC signal, heterogeneity is not solely a property of the ACS/CODH from M. thermoacetica: the ACDS from M. thermophila (0.3 spin/mol [54]) and C. hydrogenoformans (0.14 spin/mol [6]) also appear to be heterogeneous. ACS/CODH from M. thermoacetica, when expressed in E. coli (which does not naturally contain such an enzyme) is similarly heterogeneous [32]. Heterogeneity appears to be an inherent property of this group of enzymes, and does not arise by the action of an ACS-specific chaperone. Ragsdale et al. have recently reported that chelating Cu and Zn ions from populations of A-clusters followed by incubation in NiCl2 affords NiFeC spin concentrations of 0.8 spins/α [55]. If correct, these values would indicate that the heterogeneity has been essentially abolished in his samples! However, we remain cautious regarding the efficacy of this procedure. First, no such effects were observed when we performed similar procedures ([56] and unpublished results). In fact, using Mössbauer spectroscopy, we have recently found the same 30:70% heterogeneity in unactivated α subunits (i.e., before adding NiCl2) as is seen with activated subunits [47]. In these unactivated samples, the sum of Ni, Cu, and Zn concentrations were insufficient to account for the heterogeneity. Similarly, Svetlitchnyi et al. [6] reported 0.23 Zn/ α and 0.08 Cu/ α in an ACS sample from C. hydrogenoformans that yielded a NiFeC spin intensity of 0.14 spins/ α. Taken together, all of this suggests that heterogeneity arises from inherent properties of the protein rather than from heterogeneous occupancies of the proximal site of the A-cluster, though this should probably be viewed as a current controversy.

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Perhaps in the future, the best criteria for evaluating whether A-clusters within a population of α subunits are homogeneous would be to examine, in addition to the intensity of the NiFeC signal, 4.2 K Mössbauer spectra of isolated α subunits reduced with either dithionite or Ti3 citrate and then exposed to CO. In the absence of heterogeneity, all or nearly all of the Fe in such a sample would be characteristic of the Ared-CO state (S 1/2 with δ and ∆EQ values of [Fe4S4] 2 clusters) [46]. The advantage of the Mössbauer criterion is that both functional and nonfunctional heterogeneous Fe environments can be seen and quantified directly from the spectra, without significant spectral manipulation, assumptions or calculations. Understanding the origin of this heterogeneity and abolishing it will be critical for future developments in this field. Because of this problem, spectroscopic features of the A-cluster are often difficult to interpret and a number of controversies regarding the catalytic mechanism of the enzyme arise because of it.

4.3.

Reductive Activation

Methyl group transfer requires that the Aox state be reduced to a reductively activated state called Ared-act. Reduction occurs in accordance with E0 ⬃ 0.5 V compared with NHE such that only low-potential reductants such as dithionite or Ti(III) citrate are fully effective [48,57]. Most evidence favors a two-electron reduction of Aox, but a one-electron reduction has also been proposed. Bhaskar et al. [58] measured activity at various solution potentials, and concluded that a one-electron reduction of the enzyme was involved. However, Bramlett et al. [47] recently refit the Bhaskar et al. data sets by mathematical modeling and found n 1.7 ± 0.2 and E0 ⬃ 500 mV compared with NHE. In contrast, a similar experiment by Lu and Ragsdale [57] indicates n 1 and E0 ⬃ 490 mV. Spectroscopic properties generally favor a two-electron reduction, though this is rendered somewhat uncertain by the heterogeneity problem. Recent Mössbauer studies indicate that Ared-act, obtained by treating the enzyme with either Ti(III) citrate or dithionite, contains two species in a 25:75% ratio [47]. The 25% component, which would appear to reflect functional A-clusters, has parameters typical of S 0 [Fe4S4] 2 clusters. The 75% component, reflecting nonfunctional A-clusters, has parameters typical of {S 3/2:S 1/2} [Fe4S4]1 clusters. The state of Nip in the functional component cannot be observed by Mössbauer, but EPR of this state is devoid of a signal typical of Ni1 ions. The absence of Ni1 EPR could be explained by assuming that the reductively activated state involves a [Fe4S4]1 cubane spin-coupled to Nip1, and such a state has been proposed [59]. Besides being EPR-silent, reasonably strong spin-coupling would abolish magnetic hyperfine interactions in Mössbauer spectra; however, such interactions were evident for the 75% [Fe4S4]1 component of the reductively activated sample [47], thereby discounting this possibility.

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Figure 8. Mechanisms of ACS activity. Important differences include the number of electrons assumed for reductive activation (2 vs 1), the order of substrate addition (methyl group vs CO), and the requirement of an external redox pool (exclusively for the mechanism on the right). Details are given in the text.

One intriguing possibility is that both electrons localize on Nip, generating a zero-valent Ni0p state, as represented by Reaction (12) [5]: A ox {[ Fe 4 S4 ]2 Ni 2p Ni 2d}2 e A red-act {[ Fe 4 S4 ]2 Ni 0p Ni 2 d }

(12)

The achievement of such a state would be unprecedented in biology and so this hypothesis must be treated cautiously and with a healthy degree of skepticism. Nevertheless, there is substantial (though circumstantial) evidence for it [17], and we assume it in the mechanism of Figure 8, left panel. Such a state would be diamagnetic, consistent with the 25% functional portion of the A red-act α samples. Moreover, numerous Ni0 organometallic complexes have similar reactivity properties as ACS. One computational study [60] suggests that the unique ligand environment of Nip, containing three thiolates each bridged to another metal ion, may stabilize a Nip0 atom, but other studies are less supportive of this idea [59,61]. The Aox and A red-act states have been studied using Ni and Fe K-edge XAS, Ni L-edge XAS as well as soft X-ray MCD [62,63]. In the Aox state, 50% of spectral intensity was attributed to LS Ni2 and assigned to Nid, while the other 50% was attributed to HS Ni2 and assigned to Nip. When the subunit was treated with Ti(III) citrate, 30–50% of the Ni converted to another form assigned as Ni1. Whether this corresponds to a one- or two-electron activation overall is uncertain, as 50–100% of the [Fe 4S4] 2 cubanes in the A-cluster were reduced to the 1 state (or possibly to the all-ferrous state) upon treatment with Ti [62]. It was concluded that the A red-act state consists of Met. Ions Life Sci. 2, 357–416 (2007)


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2 2 1 1 0 either {[Fe 4S4] 2Ni1 p Nid } or {[Fe 4 S4 ] Nip Nid }. No evidence for Ni was reported. However, due to the heterogeneity problem, the proportion of Ni0 may have been too low (perhaps 10–20% of spectral intensity) for it to have been observed. Svetlitchnyi et al. [6] have proposed yet another electronic conformation for the Ared-act state, equivalent at the metal ions to what has been called the Aox state. They suggest that the fully oxidized state of the A-cluster contains a disulfide formed from the two cysteines that bridge Nip and Nid. This proposal is reminiscent of the D-site mechanism proposed prior to the availability of the X-ray crystal structures [48]. It could be tested by determining the X-ray crystal structure of the reductant-free form of the enzyme where the disulfide bond should be present.

4.4.

Methylated and Acetylated States

The CoFeSP protein is uniquely capable of donating the substrate methyl group to the A-cluster, with Nip the probable methyl group acceptor. The cobalamin must be reduced to the Co1 state for it to accept a methyl group from methyl tetrahydrofolate (Reaction 3). The methyl group is transferred as a cation to ACS according to Reaction (13): A red-act {[ Fe 4 S4 ]2 Ni 0p Ni 2d}CH 3 -CO3FeSP CH 3 -A ox {[ Fe 4 S4 ]2 CH 3 -Ni 2p Ni 2d}Co1FeSP

(13)

The methyl group is transferred with inversion in an SN2 type reaction [64]. The methylated state exhibits Mössbauer spectra typical of S0 [Fe4S4] 2 clusters (100% of spectral intensity) [47]. Since a methyl cation (with an even number of electrons) transfers onto the Ared-act state, the Mössbauer features of this state imply that an even number of electrons (i.e., two) is used in reductive activation. Next, CO inserts into the Ni–C bond, affording the acetyl-intermediate in Reaction (14): CH 3 -A ox {[ Fe 4 S4 ]2 CH 3 -Ni 2p Ni 2d}CO CH 3 C(O)-A ox {[ Fe 4 S4 ]2 CH 3 C(O)-Ni 2p Ni 2d}

(14)

The acetyl-bound state is EPR silent and indistinguishable from the methylated state. The last step of catalysis involves the binding of CoA and attack of the carbonyl carbon of the acetyl group by the CoA thiolate. This affords the product acetyl-CoA and regenerates the Ared-act state to complete the catalytic cycle, as Met. Ions Life Sci. 2, 357–416 (2007)

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shown in Reaction (15): CH 3 C(O)-A ox {[ Fe 4 S4 ]2 CH3 C(O)-Ni2p Ni2d} CoA A red-act {[ Fe 4 S4 ]2 Ni 0p Ni 2d}CH 3 -C(O)-CoA

4.5.

(15)

Alternative Mechanisms

There is no consensus favoring the catalytic mechanism just described, and we should mention the most popular alternative (Figure 8, right panel). This involves a one-electron reductive activation, the Ared-CO state as a catalytic intermediate, the absence of the methylated intermediate, and electron transfer steps occurring as part of the catalytic cycle [18]. Since the Ared-act state does not exhibit a Ni1 EPR signal, one must presume that this state forms in undetectably low concentrations. After binding of CO and forming the Ared-CO state, a methyl group is accepted from CH3-Co3FeSP. However, since a methyl cation is transferred, the resulting acetylated state should have a half-integer spin state, in conflict with the loss of the S 1/2 NiFeC signal upon methyl group transfer and the absence of a developing Ni3-based signal [65]. To explain this, a rapid additional oneelectron reduction from a reduced ferredoxin to ACS is presumed. The final step is the same as in the mechanism on the left in Figure 8, except that an electron is returned to the external redox pool (e.g., the ferredoxin) to complete the catalytic cycle. Bramlett et al. [47] recently reported ACS-catalyzed synthesis of acetylCoA in the absence of an external reductant, which would seem to eliminate this mechanism. It was also shown that catalysis could occur without forming the Ared-CO state.

4.6. Connection to the β Subunits In contrast to the isolated recombinant α subunit, the full ACS/CODH enzyme can synthesize acetyl-CoA using CO2 and a low-potential reductant (rather than CO) as one of the substrates. CO2 is reduced to CO at the C-cluster and then CO migrates to the A-cluster. It does this by traveling through a proteinaceous tunnel carved into the protein matrix [66]. The tunnel is not simply a tube which connects the C- and A-clusters; it has a branch point along the region between the A- and C-clusters such that the 2 C-clusters are also connected and the two β subunits are bisected [4,5]. Interestingly, the tunnel between C- and A-clusters is blocked when the α subunit is in the open conformation, and the tunnel is unobstructed when the α subunit is in the closed conformation. Tan et al. [67] also obtained evidence that the α conformation alternates between open and closed Met. Ions Life Sci. 2, 357–416 (2007)


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during catalysis. All of this implies that CO is delivered to the A-cluster at a specific step of catalysis – namely when the conformation changes from open to closed (and the tunnel opens) – and that CO is blocked from the active site at all other steps of catalysis. Tan et al. [68,69] have explored the function of the tunnel using site-directed mutants to strategically block the tunnel, either between the C- and A-clusters, or between the two C-clusters. As expected, blocking the tunnel between C- and A-clusters resulted in enzyme with normal CODH activity, but very low ACS activity. Blocking the tunnel between the two C-clusters virtually abolished CODH activity, even though all clusters appeared intact and redox-active. The only reasonable conclusion is that CO cannot access the C-cluster in these mutants, suggesting the intriguing possibility that CO (and perhaps CO2) enter and exit the enzyme through a ‘crack’ at the ββ interface. This idea requires further testing, but it is the first apparent advance in solving the mystery of how CO/CO2 enter and exit the enzyme. The A- and C-clusters also appear to communicate with each other, in that the kinetic properties of the C-cluster (kcat and KM for CO2 /CO) are affected when the A-cluster is functioning [70]. This communication may involve tunnel function [8]. In the absence of substrates required for the synthesis of acetyl-CoA (i.e., CoFeSP and CoA), CO is released during CO2 reduction assays. However, in the presence of these ‘synthase substrates’, CO is not released, but is sent directly to the A-cluster. CO at concentrations greater than ⬃100 µM was also found to inhibit the enzyme and to do so cooperatively. When the tunnel is blocked, this form of inhibition is abolished.

5.

SEQUENCE ANALYSIS AND PHYLOGENY OF THE α SUBUNIT

An alignment of twenty α subunit sequences from ACS/CODH and ACDS proteins was prepared as described for the β subunit (Appendix 3). ACS/CODHtype α subunits contain an amino-terminal domain of 30 kDa that is not present in ACDS-type subunits, as described above. ACDS-type subunits have a short carboxy-terminal extension that is rich in glutamate and lysine residues; this domain is not found in ACS/CODH-type subunits. Homologous regions of the proteins, however, are highly conserved. In addition to the four cysteine residues that coordinate the A-cluster [Fe4S4] component, the Cys-Gly-Cys motif that bridges Nip and Nid is universally conserved. The A-cluster can be viewed as being located on a wall of a crevasse. In the orientation displayed in Figure 7, one major conserved region meanders snake-like along the surface opposite/above the A-cluster. Another surface is very near the A-cluster and other conserved regions are distributed in patches around the cluster. Opposite the A-cluster on this wall are a series of conserved, negatively charged residues. One intriguing hypothesis is that these Met. Ions Life Sci. 2, 357–416 (2007)

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0.57 1.00 0.66 1.00 1.00

1.00

ANME DNA 1.00

0.98 1.00

0.93 1.00 1.00 0.81 0.76

1.00

0.1

ANME DNA

Figure 9. Phylogeny of the α subunits. This phylogeny was inferred as described for the β subunit phylogeny (Figure 5).

serve as an electrostatic guide for docking the CH3-CoFeSP prior to methyl group transfer, but this must be tested experimentally. Other conserved regions could bind coenzyme A, or facilitate the closed ↔ open conformational change. The phylogeny of α subunit proteins distinguishes separate clusters of ACS/ CODH subunits and ACDS subunits (Figure 9). Homologs in the γ -proteobacterium S. fumaroxidans and the green nonsulfur bacterium D. ethenogenes were probably acquired by horizontal gene transfer, although the donor organism cannot yet be identified. The cluster of ACDS-type subunits is similar to that of the β subunits and is consistent with vertical inheritance of these genes in the euryarchaeal lineage. The unusual placement of the M. jannaschii homologs on this tree is probably an artifact of phylogenetic inference; this placement is poorly supported by credibility values or bootstrap analysis.

6.

CORRINOID IRON–SULFUR PROTEINS

CoFeSP are 90 kDa heterodimers consisting of subunits γ and δ. The γ subunit binds a single [Fe4S4] cluster while δ binds a cobalamin coenzyme. In bacteria, a separate methyltransferase protein (MeTr) catalyzes the demethylation of methyltetrahydrofolate, transferring the methyl group to the cob(I)alamin of CoFeSP [71]. In archaea, which have sarcinapterin (and related pterin coenzymes) rather than folates, the pathway is less clear. The methyltransferase reaction could be Met. Ions Life Sci. 2, 357–416 (2007)


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catalyzed by the MtrH protein that catalyzes the analogous reaction, transferring a methyl group from N5-methyltetrahydromethanopterin to cob(I)alamin [72]. Although MtrH is considered a subunit of the methyltetrahydromethanopterin: coenzyme M methyltransferase complex, all archaea containing ACDS have MtrH homologs, including A. fulgidus, which lacks coenzyme M. Upon reductive activation to the Co1 state, CoFeSP accepts this methyl group from CH3-H4F or CH3-H4SPT (Reaction 3) and then donates it to ACS (Reaction 2). No X-ray structure has been reported for the γ or δ subunits, so little is known structurally about this protein. However, the cobalamin should be located near the surface, as a methyl group coordinated to that cobalt is (almost certainly) transferred to the Nip of the ACS α subunit. The cubane should be located within electron transfer distance to the Co ( ⬃15 Å) as this metal site serves to reductively activate the Co2 by reducing it to the Co1 state. Methyl group transfer is thought to be stimulated by protonating N5 of N5-methyl-tetrahydrofolate [73,74]. This facilitates attack by the well-known nucleophile Co1, leading to products H4F and CH3-Co3FeSP [64]. The γ subunit sequences contain four universally conserved cysteine residues near the amino-terminus that likely coordinate the [Fe4S4] cluster (Appendix 4). In the δ subunit, a set of universally conserved glutamate, aspartate, lysine, arginine and asparagine residues (Appendix 5) could bind the cobalamin coenzyme, although this protein has no recognizable sequence similarity to other cobalamin-binding proteins. Phylogenies of the γ and δ subunits are similar to each other, and consistent with those of the other subunits, clearly distinguishing clusters of ACDS and ACS/CODH proteins (Figures 10 and 11).

1.00 1.00 1.00

1.00

1.00 1.00

0.87 1.00 1.00

ANME DNA 1.00

0.98 1.00

ANME DNA ANME DNA

1.00 0.98 0.92

1.00

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Figure 10.

Phylogeny of γ subunits. Met. Ions Life Sci. 2, 357–416 (2007)

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1.00 1.00

ANME DNA ANME DNA

1.00 0.62 1.00 1.00

1.00

0.98 1.00 1.00 1.00

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0.1

Figure 11.

7.

Phylogeny of δ subunits.

ACETYL-COENZYME A DECARBONYLASE/SYNTHASES

The methanogenic archaea derive energy from the reduction of CO2, acetate (aceticlastic methanogenesis) or other single carbon compounds (methylotrophic methanogenesis) to produce methane gas. This process requires a series of conserved enzymes that are shared by all known types of methanogens, suggesting that methanogenesis arose once within the euryarchaeal lineage. Yet these different types of methanogens have evolved very different physiologies. Methanocaldococcus jannaschii is a specialist that grows solely by using H2 to reduce CO2 to methane. This physiology requires ACDS to produce acetyl-CoA for biosynthesis. Alternatively, Methanosarcina acetivorans is metabolically versatile: it can carry out both methyltrophic methanogenesis (from methanol or methylamines) and aceticlastic methanogenesis (from acetate), but cannot produce methane from formate or CO2. Thus M. acetivorans must produce acetyl-CoA or oxidize acetate depending on growth conditions. Archaeoglobus fulgidus is a sulfate reducer that probably evolved from a methanogenic ancestor and has lost the ability to grow through methanogenesis, but uses ACDS to oxidize acetate. Finally, DNA cloned from an intriguing, but uncultured group of organisms from an anaerobic methane oxidizing enrichment (ANME) encodes much of the methanogenic apparatus, including ACDS subunits [75]. These organisms may require ACDS to produce acetyl-CoA. Relatively little is known about what differentiates these enzymes from the ACS/CODH’s just discussed, but what is known is generally due to the efforts of David Grahame and his coworkers [16]. The best-studied enzyme is from Methanosarcina thermophila. These enzymes are large complexes containing five subunits (β, α, γ, δ, and ε) which are homologous to the α subunit of ACS, the Met. Ions Life Sci. 2, 357–416 (2007)


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β subunit of CODH, the γ and δ subunits of the CoFeSP and ε, a subunit with no metal centers and no known function (Table 1). The β subunits of ACDSs are homologous to the α subunits of ACS/CODHs but lack the 30 kDa amino-terminal domain (Appendix 3). This difference is responsible for the unfortunate nomenclature conflict, in that subunits are named from largest to smallest, and the lack of this 30 kDa domain causes the A-cluster-containing subunits of ACDSs to have a lower molecular mass than the C-cluster-containing subunit; hence A-clustercontaining subunits in ACDSs are called β rather than α. Note that for this review, we uniformly call the A-cluster-containing subunits α, even when referring to ACDS complexes. Interestingly, the 30 kDa N-terminal domain in ACS/CODHs abuts directly with the β subunit, requiring that the α and β subunits of ACDSs interact at a different protein:protein interface. Also, much of the tunnel that connects the two subunits in ACS/CODHs is located in this domain, suggesting a different tunnel location (or conceivably the lack of a tunnel) in ACDSs. It has been previously noted that the C-cluster-containing subunit in ACDSs contains two additional [Fe4S4] clusters (dubbed the E- and F-clusters). These clusters probably undergo redox reactions and provide an electron ‘wire’ in addition to that provided by the B- and D-clusters. If the B- and D-cluster retain their function to transfer electrons between the C-cluster and external redox agents, the E- and F-clusters might connect other active sites within the ACDS complex; note that the A-cluster and the cobalamin must both be reductively activated for catalysis, suggesting a possible role for the E- and F-clusters. The quaternary structure of the ACDSs has been proposed to be (αβγδε) 8, but this is uncertain. The 26 Å resolution structure of the (βε)2 subcomponent has been reconstructed by scanning transmission electron microscopy [76]. The 110 Å diameter globular structure reveals an internal cavity with four openings, in which the ε subunits may bridge the two β subunits. Crystals of the (βε)2 subcomponent have been reported to diffract to 4 Å resolution, but no high-resolution structure model is currently available [77]. Bhaskar, DeMoll, and Grahame have recently shown that a single ACDS serves two separate physiological functions in M. thermophila [58]. The enzyme anabolically synthesizes acetyl-CoA during chemoautotrophic growth on CO2. The same enzyme also decarbonylates acetyl-CoA as part of catabolism, namely during aceticlastic methanogenesis, a process in which energy is extracted from the degradation of acetate to CO2 and CH4 (reaction 5). Prior to this work, it was assumed that different ACDSs served different physiological functions. ACDS also catalyzes the exchange Reaction (16) where *CoA is dephosphoCoA.

CH 3 -C(O)-CoA *CoA CH 3 -C(O)-CoA* CoA

(16)

This reaction proceeds by a ping-pong mechanism by which the enzyme first reacts with acetyl-CoA, releases CoA, binds *CoA, and then releases Met. Ions Life Sci. 2, 357–416 (2007)

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acetyl-dephosphoCoA. This mechanism suggests that, after releasing CoA and before binding *CoA, ACDS is present as an acetyl intermediate. Analysis indicates that the acetyl intermediate is high-energy, such that ∆G for the hydrolysis (yielding acetate and H as products) is approximately 9 Kcal/mol. For the limited spectroscopic properties known of the metal clusters in ACDS, these clusters appear to be indistinguishable from those in ACS/CODH and have been integrated in the discussion of the A- and C-clusters discussed above. Similarly, nothing distinctive is known about the mechanism of catalysis relative to that for ACS/CODH, except the presumption that the γδ subunit homologs of CoFeSP remain bound to the αβε subunits at each step of catalysis.

8.

PHYSIOLOGICAL ROLES AND EVOLUTION OF ACETYLCOENZYME A SYNTHASE/CARBON MONOXIDE DEHYDROGENASE PROTEINS

8.1.

The Archaeal ACDSs

Molecular phylogenies of each of the ACDS subunits from the euryarchaea are similar, and the genes were probably inherited vertically throughout the euryarchaea (Figures 5, 9–11). However, these complexes function catabolically in aceticlastic methanogens and anabolically in methylotrophic or CO2-reducing methanogens. Some methanogens encode multiple ACDS proteins while others contain multiple CODH proteins or β subunits. Here we attempt to summarize the distribution and likely functions of ACDS proteins in genome sequences of euryarchaea (Appendix 6).

8.1.1.

Methanobacteriales

Methanothermobacter thermautotrophicus was isolated from an anaerobic sludge digester where it probably grew by methanogenesis using CO2 and H2, produced by fermenting bacteria in coculture. The genome contains a cluster of seven genes that includes the αβγδε subunits of ACDS, a nickel insertion protein (CooC) and an iron–sulfur protein (CooF). Adjacent to this cluster are genes involved in cobalt uptake and cobalamin biosynthesis, required by the CoFeSP. These cells incorporate exogenously supplied radiolabeled CO into acetyl-CoA [78], indicating the function of the ACDS in this organism.

8.1.2.

Methanomicrobiales

Similar to M. thermautotrophicus, Methanospirillum hungateii is a CO2-reducing methanogen that is commonly isolated from anaerobic digestors. Cells readily Met. Ions Life Sci. 2, 357–416 (2007)


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form syntrophic cocultures with anaerobic fatty-acid-oxidizing bacteria by consuming H2, CO2, and formate fermentation products. The complete genome sequence of M. hungateii encodes the five subunits of an ACDS complex and two copies of CooC in a single gene cluster. A second cluster encodes CooF, a bacterial-type β subunit homolog, and a NADH-quinone oxidoreductase.

8.1.3.

Methanococcales

These marine methanogens use hydrogen or formate as electron donors to reduce CO2. They are autolithotrophs and cannot use acetate or methyl compounds as sole carbon sources. Therefore they rely on an ACDS enzyme to produce acetylCoA for biosynthesis [79]. Methanococcus maripaludis has a single gene cluster encoding all five subunits of an ACDS enzyme, along with two copies of cooC and one cooF gene. A similar ACDS enzyme was previously identified in the related species Methanococcus vannielii [80]. Methanocaldococcus jannaschii encodes the five subunits of a complete ACDS enzyme in two gene clusters. The first cluster encodes α, β, and ε subunits in addition to CooF and a paralogous α subunit. One α subunit contains an aminoterminal domain that is characteristic of ACS/CODH proteins; but the remainder of the protein sequence is most similar to its M. jannaschii paralog. This suggests the gene may be chimeric. The second cluster encodes γ and δ subunits of the CoFeSP complex. Another β subunit, which is related to bacterial CODH-like proteins, is missing two cysteine residues involved in C- and D-cluster formation. The gene lies in a cluster that includes paralogs of the coenzyme F420-reducing hydrogenase complex genes. This system could be analogous to the CODH/ hydrogenase complex found in the bacterium Rhodospirillum rubrum, which is described in Section 8.2.2. Alternatively, M. jannaschii could couple the oxidation of CO (E0 0.52 V for CO2) with F420 reduction (E0 0.35 V).

8.1.4.

Methanopyrales

Methanopyrus kandleri is a highly diverged marine autolithotrophic methanogen that specifically carries out methanogenesis through CO2 reduction using H2. A single gene cluster encodes subunits of the ACDS complex and a second paralog of the β subunit. One of these proteins lacks cysteine residues essential for forming the B and D [Fe4S4] centers and is missing two other conserved cysteines in the C-cluster that are not ligands for the [Fe3S4] subsite. The function of this aberrant protein is unknown and it is highly unlikely that it catalyzes Reaction (1). This ACDS gene cluster also contains cooF and cooC genes and is associated with the pyruvate:ferredoxin oxidoreductase (PFOR) gene cluster, which encodes an enzyme catalyzing pyruvate synthesis through the reductive carboxylation of acetyl-CoA. Alternatively, the combination of ACS, CODH, and PFOR Met. Ions Life Sci. 2, 357–416 (2007)

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enzymes forms an efficient hydrogenase in the presence of CO and pyruvate [81]. M. kandleri also encodes a third copy of the β subunit belonging to the bacterial CODH-like group. As in M. jannaschii, this gene is associated with paralogs of the coenzyme F420-reducing hydrogenase genes.

8.1.5.

Methanosarcinales

Members of the Methanosarcinales are some of the most metabolically versatile methanogens. Their relatively large genomes result from duplications and extensive gene acquisition by horizontal gene transfer. The ACDS complex from Methanosarcina thermophila is the best-studied example of the archaeal-type enzyme [82]. Methanosarcina thermophila is an acetotrophic methanogen that was originally isolated from an anaerobic digester. Cells carry out methanogenesis using either acetate or one-carbon substrates. Despite strain heterogeneity, M. thermophila cells appear to use a single ACDS complex for both acetate cleavage (aceticlastic methanogenesis) and acetyl-CoA synthesis (for biosynthetic purposes) [83,84]. Genes encoding the five subunits are found in a single operon, cotranscribed during growth on acetate [85]. Between genes encoding the αβγ and δε subunits of ACDS there is a cooC gene. Transcription of these genes is highest during growth on acetate compared with growth on methanol, indicating that tight regulation of gene expression rather than the differential expression of isozymes (subfunctionalization) is the primary mode of adaptation for these cells [86]. However, we cannot rule out the presence of additional subunit homologs until a complete genome sequence of M. thermophila becomes available. The marine methanogen Methanosarcina acetivorans carries out methylotrophic methanogenesis (from methanol or methylamines) and aceticlastic methanogenesis (from acetate), but cannot produce methane from formate or CO2. The six-gene operon encoding an ACDS complex and CooC has recently duplicated in this organism: the individual protein sequences are 82–100% identical [87]. Three more homologs of the β subunit are scattered throughout the genome. One of these paralogs is identical to a β subunit from an ACDS complex operon. The other two paralogs encode bacterial-type CODH proteins. One is associated with an iron–sulfur electron transport protein. The second paralog belongs to the CODH-like family of proteins with modified D-cluster and C-cluster residues. Recently M. acetivorans was grown anaerobically on CO under conditions that inhibited methanogenesis and led to the production of formate or acetate [88]. The uncharacterized bacterial-type CODH proteins could facilitate this carboxydotrophic growth. Like M. acetivorans, Methanosarcina mazeii cells carry out methylotrophic and aceticlastic methanogenesis, but can also grow on H2 and CO2. Strains of this organism have been isolated from natural anaerobic environments as well Met. Ions Life Sci. 2, 357–416 (2007)


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as anaerobic sewage digestors. The ACDS subunit complement of M. mazeii is similar to that of M. acetivorans with duplicate gene clusters. Two orthologs of the bacterial-type CODH proteins found in M. acetivorans are also present in M. mazeii. Thus the common ancestor of these two methanosarcinales probably had a similar complement of ACDSs and CODHs. DNA microarray experiments using M. mazeii showed the expected high levels of ACDS gene expression during growth on acetate as well as enhanced expression of iron and cobalt ion transporters [89]. Increased expression of both ACDS operons was reported, suggesting that gene duplication occurred to enhance expression levels, not to evolve new functions or to subfunctionalize the ACDSs. However, it is not clear that the hybridization methods could distinguish between these very similar sequences. No significant change in expression of the bacterialtype CODH genes was reported for growth on acetate versus methanol. Methanococcoides burtonii is a methylotrophic methanogen that cannot grow on acetate. Although its genome sequence is incomplete, it appears to have one ACDS complex encoded by a single gene cluster. In contrast, Methanosphaera stadtmanae grows by methanogenesis through the H2-mediated reduction of methanol. This species requires acetate for growth and has no homolog of any ACDS subunit in its genome [90]. Nor does M. statdtmanae have orthologs of cooC or cooF.

8.1.6.

Archaeoglobales

Archaeoglobus fulgidus is a sulfate reducer that probably evolved from a methanogenic lineage by acquiring genes for dissimilatory sulfate reduction and fatty acid degradation while losing the capacity to grow through methanogenesis [91]. This heterotrophic archaeon has an ACDS that functions in acetate oxidation [92,93]. The five genes are arranged in one cluster containing γδα (with cooC) and two paralogous clusters of βε genes. Only one type of ε subunit co-purified with the ACDS complex [93]. The gene cluster containing the unexpressed ε subunit gene also encodes an aberrant β subunit with unusual Asn and Gln ligands to the B- and D-clusters. Therefore, we expect these genes are either not expressed or have a different function. An additional β subunit gene is not associated with any genes of known function and encodes a CODH-like protein with modified C- and D-cluster residues.

8.2. ACS/CODH Genes in Bacteria 8.2.1.

Clostridia

The clostridia are anaerobic heterotrophs that can oxidize or ferment a wide variety of organic substrates [94]. Acetogenesis, the formation of acetyl-CoA by Met. Ions Life Sci. 2, 357–416 (2007)

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CO2 fixation, was first elaborated in Clostridium thermoaceticum (now Moorella thermoacetica) [95] and the Wood-Ljungdahl pathway was first discovered in this organism. Because this pathway requires an ACS/CODH enzyme to catalyze acetyl-CoA biosynthesis, the ACS/CODH protein from M. thermoacetica has been well studied and serves as a model system [96]. Besides M. thermoacetica, clostridial genomes that encode a complete ACS/ CODH complex include Alkaliphilus metalliredigenes, Carboxydothermus hydrogenoformans, Clostridium difficile and Desulfitobacterium hafniense. ACS/CODH genes from these bacteria often cluster with cooC genes and genes encoding other proteins in the Wood–Ljungdahl pathway. While most clostridia do not have an ACS/CODH complex (or use the Wood– Ljungdahl pathway), numerous clostridial genomes encode homologs of the single subunit CODH protein. Pathogenic clostridia (Clostridium botulinum, Clostridium difficile, and Clostridium tetani) contain one or more copies of the CODH gene, as do environmental and solvent-producing clostridia such as Clostridium acetobutylicum, M. thermoacetica, and Ruminococcus albus. These bacteria are phylogenetically scattered throughout the clostridia. Therefore we may infer that the ancestor of the clostridia encoded a similar protein and that the CODH enzyme is quite old. Carboxydothermus hydrogenoformans is a hydrogen-producing, carbon monoxide-oxidizing bacterium that can use CO as a primary carbon source. The genome sequence of the microbe revealed one ACS/CODH and four CODH enzymes [97]. The bacterium uses the ACS/CODH complex (called CODH-III) in the Wood–Ljungdahl pathway to fix CO. One CODH enzyme (CODH-I) is genetically linked to a hydrogenase gene cluster, suggesting that it forms the CO-oxidizing:H2-evolving complex that was previously isolated from C. hydrogenoformans [98]. This complex is similar to the well-studied Rhodospirillum rubrum complex that is described in Section 8.2.2. Two of the remaining CODH proteins (CODH-II and CODI-IV) are associated with CooF proteins and have been proposed to support oxidative stress response or anabolic electron transfer processes [97]. The fifth homolog (CODH-V) represents the CODH-like group of bacterial and archaeal proteins with modified C- and D-cluster residues and no known function. Desulfitobacterium hafniense, another member of the Peptococcaceae family, has a similar complement of ACS/CODH and CODH proteins — all are most closely related to orthologs in C. hydrogenoformans. Dehalococcoides ethenogenes is a tetrachloroethene-dechlorinating member of the green nonsulfur bacteria that grows on acetate. Its genome encodes homologs of the α, γ and δ subunits of an ACS/CODH complex (all in a gene cluster with other Wood–Ljungdahl pathway genes), yet there is no apparent homolog of the β subunit in this genome [99]. The genome is also missing methylene-tetrahydrofolate reductase and methyltetrahydrofolate:cob(I)alamin methyltransferase genes. It is not clear how any of the identified components of this Met. Ions Life Sci. 2, 357–416 (2007)


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pathway could function without these essential proteins. These genes were almost certainly acquired by horizontal gene transfer, but we do not know whether the missing genes were not transferred or lost subsequent to the transfer event.

8.2.2.

Proteobacteria

Only one of the many sequenced proteobacterial genomes encodes homologs of all four ACS/CODH/CoFeSP subunits. This strict anaerobe, Syntrophobacter fumaroxidans, grows syntrophically with hydrogen- and formate-consuming methanogens in a propionate-oxidizing coculture [100]. Originally isolated from an upflow anaerobic sludge bed, S. fumaroxidans cells oxidize propionate using the methylmalonyl-CoA pathway, releasing hydrogen and formate that is consumed by methanogens. This organism can also be grown in pure culture by fumarate fermentation. Enzyme activities of the Wood–Ljungdahl pathway (including CODH) were found in extracts of fumarate-grown cells. This pathway was suggested to be reversible, functioning as an anaplerotic reaction in propionate catabolism, and an aceticlastic reaction during fumarate fermentation [101]. S. fumaroxidans was grown in pure culture on propionate and sulfate, but could not be grown on acetate and sulfate. Other syntrophically grown bacteria can grow on acetate in coculture with a methanogen using the Wood–Ljungdahl pathway [102,103]. While there is ample opportunity for gene acquisition by horizontal gene transfer in these consortia, the phylogeny of the ACS/CODH genes indicates the S. fumaroxidans genes were recruited from the clostridia, not from methanogens. Its genome contains a second CODH gene that is associated with other oxidoreductase genes. Rhodospirillum rubrum is a purple nonsulfur photosynthetic bacterium that oxidizes CO. The CooA heme protein senses redox state and CO, activating transcription of CODH during anaerobic photosynthetic growth [104]. CO oxidation by CODH is coupled to hydrogenase activity by the ferredoxin-like electron transfer protein CooF [105,106]. Tight regulation of CODH expression is probably important in this organism because it can also grow heterotrophically as a facultative aerobe. A similar set of genes is found in the genome sequence of another α-proteobacterium, Rhodopseudomonas palustris str. BisB18, although these genes are missing from sequences of five other R. palustris strains. This distribution may have arisen from a recent gene transfer event. The sulfatereducing δ -proteobacterium Desulfovibrio vulgaris also has homologous CODH and hydrogenase genes that were probably acquired by horizontal gene transfer. Several δ -proteobacterial genomes have homologs of the R. rubrum CODH, but are missing the associated hydrogenase genes. These organisms include Desulfovibrio desulfuricans, Geobacter metallireducens, Geobacter sulfurreducens, and Pelobacter carbinolicus. Similarly, the green sulfur bacterium Chlorobium Met. Ions Life Sci. 2, 357–416 (2007)

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phaeobacteroides DSM 266T has an unassociated CODH gene, while other Chlorobium species have no homologs. CODH genes in these genomes are usually associated with cooC and cooF genes. These organisms may couple CO oxidation to unknown reductive processes.

9. ORIGINS AND EVOLUTION OF ACDS, ACS/CODH, AND CODH COMPLEXES In the chemoautotrophic (or Fe/S world) scenario for the origin of life [107–109], the first autocatalytic metabolic pathways evolved through inorganic reactions on mineral surfaces. According to this hypothesis, the Earth was cooling but still hot ⬃3.4 billion years ago and the only compounds available were simple molecules such as CO2, N2, H2S, etc. The first event in this process was the reduction of CO2 on a Ni–Fe–S surface to various activated forms, e.g., {CH3–Ni,Ni– CO,CH3C(O)–Ni}. These reactions, which are analogous to those found in the modern reductive acetyl-CoA pathway, have been shown to occur in vitro [110]. In the transition from surface metabolism to modern nucleic acid or protein-catalyzed reactions, heterogeneous metal catalysts became replaced by protein-based equivalents as new ligands coordinated aquated transition metals and reconstituted Ni–Fe–S surface-like scaffolds within protein active sites. Reconstructing the evolutionary events which have resulted in modern ACDS, ACS/CODH, and CODH enzymes is obviously speculative, but interesting nonetheless. It seems reasonable to assume that CODH and ACS components evolved independently and eventually merged. For example, CODH could have evolved from an ancestral protein containing 2–3 [Fe4S4] clusters (analogous to Nideficient CODH). Once one of these clusters evolved the means to install a Ni ion (perhaps via mechanism B of Figure 4), this primitive CODH could have reduced CO2 to CO or have coupled the oxidation of CO to the reduction of another oxidoreductase, e.g., a membrane-bound hydrogenase, as a way to extract energy from a CO-rich environment. This same ancestor might have been shared by the hybrid cluster protein, which contains a modified [Fe4S4] center reminiscent of the C-cluster [41]. CoFeSP may have evolved from the merger of another [Fe4S4] containing protein with a cobalamin-containing protein able to transfer methyl groups. ACS might have evolved from an ancestral protein containing a single [Fe4S4] cluster. Once the critical C595-G596-C597 sequence arose, two Ni ions could bind, affording a primitive A-cluster which allowed the protein to catalyze the synthesis of acetyl-CoA using CO (but not CO2) as one of the substrates, and the methyl group from CoFeSP as another. If the organism housing this enzyme existed in a CO-rich environment, this ability would have provided a new means to synthesize acetyl-CoA. The eventual merger of ACS and CODH may have been facilitated by a decline in environmental CO and an increase in CO2. Such a merger would have allowed ACS to continue to synthesize acetyl-CoA using CO2 Met. Ions Life Sci. 2, 357–416 (2007)


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as a substrate. In some lineages, the CoFeSP heterodimer remained independent of ACS/CODH while in others, it merged to generate ACDS complexes. This model does not fully connect the CO2-based surface metabolism predicted by Wächtershäuser [107] to the modern CODH, ACS/CODH or ACDS enzymes, as it assumes a CO-rich environment. Neither does the model predict the direction of the CODH reaction in the ancestor nor explain the enzyme’s physiological role in a primitive ancestor. However, this model requires fewer instances of gene loss or horizontal gene transfer to explain the current distribution of homologs in anaerobic microbes than alternatives. It also posits simple ancestral [Fe4S4] proteins that acquired catalytic functions through mutation and uptake of Ni ions, followed by the merger of subunits to generate the range of function and subunit associations found in the group of extant enzymes. An alternative scenario assumes that the primitive enzyme was an ACDS similar to those found in modern methanogens. After formation of distinct archaeal and bacterial branches, the enzyme in the archaeal branch evolved predominantly by vertical gene transfer and its components became tightly integrated in the cell’s metabolism. Due to these constraints, modern ACDSs are not easily acquired by lateral gene transfer. In bacteria, the enzyme became modular and more amenable to horizontal gene transfer, especially as these organisms become closely associated with diverse anaerobic bacteria. In some lineages exposed to CO-rich environments, ACS components were lost and cells grew by oxidizing CO to CO2 in conjunction with a membrane-bound hydrogenase. This model attractively proposes an enzyme of clear utility in the universal ancestor: acetyl-CoA produced by this system can be the precursor for most modern metabolites. However, this model assumes that a complicated protein evolved in an early, primitive ancestor, and it requires the parallel loss of these genes in numerous phylogenetic lineages to explain the modern gene distribution. In any event, it is clear that the physiological roles of these enzymes have changed as the organisms evolved to their current configurations, and that throughout this process, they have played a dominant role in extracting energy and/or cell carbon from the environment [19]. The evolutionary history of these enzymes will become clearer as more sequences become available and as more is learned about the metabolisms of organisms containing these sequences. Determining the structures of an ACDS and CoFeSP as well as a more complete characterization of the mechanistic enzymology of ACDSs vis-à-vis the ACS/CODHs will also be of paramount importance in establishing the role that these remarkable enzymes played in the history of life on this planet and the role they continue to play in the global carbon cycle.

ACKNOWLEDGMENTS New work on this review was funded in part by a grant from The Welch Foundation (F-1576), the National Science Foundation (MCB-0425983), and the National Met. Ions Life Sci. 2, 357–416 (2007)

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Institutes of Health (GM46441). We thank Dr Lisa Perez and the Laboratory of Molecular Simulation at Texas A&M University for preparing Figures 1 and 7.

ABBREVIATIONS ACDS ACS/CODH ANME CoA CODH CODHCh CODHMt CODHRr CODHRr* CoFeSP ENDOR H4F H4SPT MCD PFOR XAS

acetyl-CoA decarbonylase/synthase acetyl-CoA synthase/carbon monoxide dehydrogenase anaerobic methane-oxidizing enrichment coenzyme A carbon monoxide dehydrogenase carbon monoxide dehydrogenase from Carboxydothermus hydrogenoformans carbon monoxide dehydrogenase from Moorella thermoacetica carbon monoxide dehydrogenase from Rhodospirillum rubrum carbon monoxide dehydrogenase from Rhodospirillum rubrum, inactive precursor form devoid of Ni corrinoid iron–sulfur protein electron nuclear double resonance tetrahydrofolate tetrahydrosarcinopterin magnetic circular dichroism pyruvate:ferredoxin oxidoreductase X-ray absorption spectroscopy

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APPENDICES Appendix 1. Spectroscopic Properties of the C*-cluster All Fe species in CODHRr* are observed proportionately by Mössbauer spectroscopy; thus, 40% of spectral intensity would be expected to arise from the B-cluster, 40% from the C*-cluster and 20% from the D-cluster. In the oxidized state, 85–90% of the Fe in CODHRr* is diamagnetic with isomer shift and quadrupole splitting parameters (δ and ∆ EQ) typical of [Fe4S4] 2 clusters, while 10– 15% of the Fe appeared to arise from [Fe3S4]1 clusters formed from the oxidative decomposition of Fe4S4 clusters [23]. In the fully reduced state, ⬃100% of the Fe was magnetic in the absence of an applied field, with 60% typical of S 1/2 [Fe4S4]1 clusters and 40% typical of S 3/2 [Fe4S4]1 clusters. The former was associated with a gave 1.94 signal, allowing this portion of the Fe to be assigned to the Bred cluster (and perhaps, in hindsight, the Dred cluster: note that 40% for the B-cluster 20% for the D-cluster equals the observed 60% spectral absorption). The latter was associated with EPR features between g 6–3, as is typical of S 3/2 [Fe4S4]1 clusters. In the reduced samples of CODHRr*, there was no evidence of [Fe3S4] clusters; such clusters would have been detectable at the 10–15% level. Nor was there evidence for Feu (i.e., a HS ferrous ion). Both Mössbauer and MCD spectra are difficult to analyze, but analysis of MCD spectra is particularly difficult because only centers that possess particular optical and magnetic properties can be observed. Even if such centers are observed, spectral intensities are not directly proportional to the concentration of the center in the sample. Regarding the particular case at hand, numerous spectral subtractions, re-scalings, and assumptions were required to isolate the MCD contribution of the C*red cluster [20]. Although each manipulation seems reasonable, collectively they add uncertainty to the resulting spectral features assigned to this cluster. Moreover, given the unique structure proposed (an Fe3S4 cluster spin-coupled to Feu), the MCD of standard [Fe3S4]1/0 clusters may not be comparable. This is so because spectra depend (exquisitely) on the magnetic properties of the system. For example, the oxidized C* cluster exhibits no MCD because it is diamagnetic, whereas oxidized [Fe3S4]1 clusters exhibit MCD because they are S 1/2 [113]. This inconsistency is not viewed as a problem because spin-coupling to Feu could render the system diamagnetic (e.g., if the unique Fe was LS ferric and antiferromagnetically coupled to the S 1/2 of the [Fe3S4]1 cluster). On the other hand, the MCD of the reduced C* cluster is compared with that of a standard [Fe3S4] 0 cluster, and this comparison forms the basis for the proposal that C* has a [Fe3S4] core structure. Given that Feu is HS ferrous under the reducing conditions where C*red would be present, spin-coupling to such an ion would change the fundamental magnetic properties of the system such that the MCD of C*red would be unrelated to that of a standard [Fe3S4] 0 cluster [113]. Met. Ions Life Sci. 2, 357–416 (2007)

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Appendix 2. Alignment of β Subunit Protein Sequences R .r u b r u m M o. t h e r m o 2 C .p h a e o b C .d i f f i c i l 2 M .j a n n 2 M .k a n d l 3 D .h a f n i e n 5 C .h y d r o 5 M s. a c e t 4 S .f u m a r o x 2 C .h y d r o 2 D .h a f n i e n 2 A .m e t a l l 1 C .h y d r o 1 C .h y d r o 3 D .h a f n i e n 3 D .h a l f n i e n 1 C .h y d r o 4 A .m e t a l l 2 C .d i f f i c i l 1 G .s u l f u r M .h u t c h i n 1 S .f u m a r o x 1 M s. a c e t 3 A .f u l g i d 3 M o. t h e r m 1 M c. j a n n 1 ANME1 A .f u l g i d 2 A .f u l g i d 1 M .h u t c h i n 2 M c. m a r i p a l M .t h e r m a u t M .k a n d 1 M .b u r t o n i i M s. t h e r m o M s. a c e t i v 2 M s. a c e t i v 5 ANME2 M .k a n d 2

1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

M - T H --------------------------------------HDCAHCSSDACATEMLNLAEA------NS I E T AWHRYEKQQ-M - - - ------K-YSYPIL-----------------NAEMPGREDVLRLTPNPASKDLLEYLHQ------EG VETWLDRYEAQQ-M A E E RIYRYDDELTGSRNAPQYQPGMSLKERIKLVHRLNYSKEEVIKHTANKAVAEMVEHMDK------EA I S N TFDRFAQQH-M - - - --------------------------------------ACNTCKMCESADKKLESFVAS------KD V E T AFHRTEDQK-M - - - ----------------------------------------KNCIAAIPEVKEMVEKAKL------KG IETPHTRFPNQF-M - - - ------------------------------------EEKRSSCPYADEAVCELVEHAKE-LNEEIPE IETPHIRWPVQF-M - - - --------------------------------SETVLEKTEGRVSYHDSVEEMLVRIRE------DG M S N VFDRYQAQE-- - - - ---------------------------------------MATKTSIHPSVNELYQRLAE------DQ L S N CFDRFDPQE-M - - - ---------------------------------------DKERISYHESVQKMYERIKK------DN M T N VWDRYEAQGIG M S Q E E-----K-KEKQLSEV---------------KKSKP--LDPKAATTCESTIQMLRRAAA------DGAETA FNRAAEMK-M - - - ---------------------------------------SNWKNSVDPAVDYLLPIAKK------AG I E T AWDRYEAMQ-M - - - ----------------------------------------SLRFSIDEAVQKLLPVAEK------DK I Q T VWDRHKDQQ-M - - - ------------------------------------------GNCHNLSNQCLINKAED------EN I D L VWHRYEKML-M - - - --------------------------------------AKQNLKSTDRAVQQMLDKAKR------EG I Q T VWDRYEAMK-M - - - ---------------------------------------DKSKLSVDPVIPNLYRKARE------EG I S T VFDRYEAQQ-- - - - ---------------------------------------MLNVRSIDPAAQDLIKKAHK------DG I E T VWDRYDKQQ-M P R Y R-----D-LTHTSRPS---------------NAPRVVEPKNPLRTTDPGTIEMLKVAQE------NK L E T VFDRFVAQQ-M P R F R-----D-LEHTSKPS---------------KADRVWEPKNRKRTIDPAALEMLEKAEK------DG V K T AFDRFVEMQ-M - - - --------------------------------------SEEKMVSLNTIDDQLIKKAAD------EG I E T MWDRKKAMK-M - - - ---------------------------------------DEKMLTIDLNSQKMIAKARE------EG V E T MYDRKEGFK-M D Q - ------------------------------------ARNGHDSRSIDPAAKEMLRIADR------EGYATI WERYEQQQ-M T E - ----------------------------------QDIRIEAERRASDRTDQEIIRKTLD------EG V Q T VWDRFALQQ-M N D - ---------------------------------------TYAGRTITRDGQLLLEKAEK------DR V E T VWDRFENQL-M G K - -----------------------------------EMKQTAFEKSIDSASQIMLQKAED------EG I E T AWDRYEQQL-M - - - --------------------------------------KIEGKVSEHESINMMYERVSK------EG VTNIVDRFNAQE-M P R F R-----D-LSHNCRPS---------------EAPRVMEPKNRDRTVDPAVLEMLVKSKD------DK V I T AFDRFVAQQ-M V M G NNVEMDIKKL-LTPLVKMKNANISMSIKFG-EEE----------------EEEWEPMGPTPMPKIPT L R H -WDFKLLERYP M - - - ----EEIKKGGIFTIPELQNVRINIGEIIEE-EE-----------------E-WEPMGPHPMPRIAT L R D -WDFKLLNRYK M - - - ---VLEFGKGA-FVVDDLRNVTIKIGEIAEE-EE-----------------E-WAPMGPTPMPGIAT L R D -WDFFLLKRYK M - - - ----FELKKGA-LFVDEMKNVSIRIGKVVEE-EE-----------------EVWEEAGPTPKPGILE L R K -WDHKLLERYE M - N R KNIHIRLSEAK-SSTGSIKDLDIKIGKIKGE-QW-----------------E--EKEGPTPFPDHTA L R N -WDRMLLTRYK M A Y K KM----DKKL-DDDFWKTKNLSLSIGEVSVGKPK----------------QELDEPMGPTPKPSVTD L R S -WDFKLLKRYP M I D V APESKKAKDL-KGDFWDAKNIQISIGEIITE-EK----------------PPEEEVKGPKPRPHVTD L R S -WDMKLLERYE M - - - SPFEL-----------EFEGLKVQIGSIEGF-EP--------------------RGEGPLPCPTVSD L A D -WDRKLFARYR M - - - ----SELTTGR-FSISDLDNVQITINNIVGA-IE---------KQSDDIDVE----MGPTVKPGVSS L R D -WDHNILDRYN M - - - ----AKLTTGS-FSIEDLESVQITINNIVGA-AK---------EAAEKAEEE-LGPMGPTPFPTAAT V R D -WSFTLFDRYE M - - - ----SKLTTGS-FSIEDLESVQITINNIVGA-AK---------EAAEEKAKE-LGPMGPTAMAGLASYRS- WNLLLLDRYE M - - - ----SKLTTGS-FSIEDLESVQITINNIVGA-AK---------EAAEEKAKE-LEKAGPTLFPGLESYRDD WNFKLLDRYE M - - - ------IKDGR-FILDGMENVVINIGKVTTE-EE----------LAE----EEWEPMGPTPKPGILS L R N -WDHRLLKDFP M - - - ---GI-----------EWEGVKVEIGELVVE-GDSE---------------------SEMEGPTRRE L L P -WDRTLASVYD

R .r u b r u m M o. t h e r m o 2 C .p h a e o b C .d i f f i c i l 2 M .j a n n 2 M .k a n d l 3 D .h a f n i e n 5 C .h y d r o 5 M s. a c e t 4 S .f u m a r o x 2 C .h y d r o 2 D .h a f n i e n 2 A .m e t a l l 1 C .h y d r o 1 C .h y d r o 3 D .h a f n i e n 3 D .h a l f n i e n 1 C .h y d r o 4 A .m e t a l l 2 C .d i f f i c i l 1 G .s u l f u r M .h u t c h i n 1 S .f u m a r o x 1 M s. a c e t 3 A .f u l g i d 3 M o. t h e r m 1 M c. j a n n 1 ANME1 A .f u l g i d 2 A .f u l g i d 1 M .h u t c h i n 2 M c. m a r i p a l M .t h e r m a u t M .k a n d 1 M .b u r t o n i i M s. t h e r m o M s. a c e t i v 2 M s. a c e t i v 5 ANME2 M .k a n d 2

39 51 78 37 35 44 43 35 38 55 36 35 33 37 36 35 57 57 37 36 41 43 38 42 37 57 67 59 59 59 63 64 67 50 63 66 66 67 60 46

- - - -PQ CGFGSAGLCC RIC LK GP CR IDPF-GEGPKY GVC G ADRDTIVARHLVRMIAA GT A A HS E HGRHIALAMQHISQGELHDYS - - - - PM CGYGLRGLCC RMC QW GP CR LDNK----RQR GIC G RDLSTVIMANLVRSLVA GL A A HG R HAHEVILTIMAAAEGKA-NLP - - - -PQ CGYGLTGACC AFC SY GP CR VTEK----TLYSV C G KDVDLIVAGNALRRLAS GM A A HG A HAREVFIALKAAAEGSA-PIP - - - - VKCGFGLQGVCC RLC SN GP CR VTPK----SPR GIC G ADADTIVARNFLRAVAS GA ACYLHVVENTAKN LKNVGITKG---V - - - -PK CPYGLKGVYC ILC AN GP CR ITEK----TPY GVC G ATADVIVARNLCRAVAA GTSCYVHC AENAARALLSAGKGEG-SYE - - - -PK CPYGKQGVWC NIC SN GP CR ITEK----TPR GVC G ATADVIVARNFLRHVAA GA ACYVHCLENAARA LKSVADEES-PYE - - - K IR CKFCLQGLTC QQC SQ GP CR INEK-GE-QDR GVC G IGPDAMAMRKLLLQNIM GA GTYSH HAYEAFRTLKATGEGKT-PFK - - - K IRCNYCELGVSC QLC SN GP CR INEKVG--ATL GVC G INADGMAMRYMLLRNVM GTSTYTY HAYEAYKTLKMTALGNT-PFT G V P D RRCTFCMAGARC DLC SN GP CRSDAAKD---KR GVC G ITADGMAMRMMLLRNVM GASTYHY HTDQTIRTLRETAKGKT-PYS - - - - P- CPIGAESSCC KHC FM GP CR LNPR-DPYSRV GVC G ATIDTIAARNFSRMVAS GG A A HT D HGMGMLDVFREVVRGNIKGYK - - - - PQ CGFGELGVCC RIC WK GP CR IDPF-GNGPQR GVC G ADAHTIVARNLIRMIAA GA A A HS E HGRHIALTLLEVGEGHAPAYR - - - - PQ CGFGKMGICC RIC WK GP CR VDPF-GKGAQR GIC G ADAHTIVARNLIRGIAA GA A S HS D HGRHIALTMREVGKGNASAYT - - - - PL CGFGEKGICC RIC MK GP CR IDPF-GNGPQE GIC G ANADTIVARNLVRMIAA GT A A HS S HGKEIAKTLMQVGKKTSKSYA - - - -PQ CGFGETGLCC RHC LQ GP CR INPF-GDEPKV GIC G ATAEVIVARGLDRSIAA GA A G HS G HAKHLAHTLKKAVQGKAASYM - - - -PQ CGFGLTGLCC RHC VQ GP CR IDPF-GEGPQA GIC G ATAEVITARNLLRQVTA GA A A HV D HAYDVLEVLEQIAQGTE-SYS - - - - PQ CGFGSLGVCC RHC IQ GP CR IDPF-GQGPDK GIC G ATADVIVARNLLRQVAA GA A A HV D HAYDAVEALELAAQGKI-DYP - - - -PQ CGFGYKGICC RIC LA GP CR VKAE-DGPASR GIC G ATAYTIVSRNLVRMIAG GA A S HS E HARHVLHTAHELVEGKTKDYE - - - - PQ CQFGYKGLCC RFC LQ GP CR LP-N-DDPSKK GIC G ASAWTIAARSVGTLILT GA A A HN E HARHIAHALKELAEGKAPDYK - - - - AP CGFGEKGVCC RIC GM GP CR VSPVAGKGAQK GIC G ATADTIVARNFARMVAA GT A S HS D HARDIAHVLHMASRD--GNYQ - - - - AQCGFGLQGVCC RIC GM GP CR ISPK----TPR GLC G ADEHTIVGRNFARMVAG GT A A HS D HARDIAHTLALADPN--GNYK - - - -PQ CSYGQLGTCC RIC SM GP CR IDPF-GDGPTR GVC G ATADTMVARNLARMAAV GS S S HS D HGRKVALLLKAVANGSNTDYH - - - - PQ CGFGQLGVCC NNC AM GP CR IDPF-GGKTSL GVC G ANVDTIVARNLLDDLSV GA A A HS D HGREVVQVLLETAEGHGQGYQ - - - - PQ CGYCEMGLSC RIC VM GP CR IDPF-GEGPQK GVC G ADADIIVARNLCRMIAA GA A S HS D HGRDLIEVLDEVAKGKAAGYR - - - -PQ CSFGQLGICC RNC NM GP CR IDPF-GEGTEK GIC G ATADIIVARNLLRMIAA GA A A HS D HARDAVLTFKKMSEGKAGSYR - - - K GR CPFCEKGLSC QLC SM GP CR ISKDK----PT GAC G IDAAGMVVRNFTHKNML GTEAYTY HAIEAAKTLKATAEGKT-IYE - - - - PQ CKIGYEGICC RFC MA GP CR IKATDGPGS-R GIC G ASAWTIVARNVGLMILT GA A A HC E HGNHIAHALVEMAEGKAPDYS P F Y MPI CD-----LCC -LC TF GK CD LS-----RGKK GAC G LNIKAQQARIVLIACCI GA A C HA G HSRHLVHHLIETLGR---DYP P F Y APL CD-----MCC -LC TY GK CD LT-----GNKK GAC G IRMDAQQGRIVLLACLV GC S A HC G HGRHLLHDMIKKLGP---DVP P F Y A PA CD-----MCC -LC TM GK CD LT-----GNKR GAC G IDLAAQTGRIVTIACSI GV S A HT G HARHMLHDIEHMTGKKLSEIP P F Y A PMQD-----F CN-LC TM GP CD LS-----MNKR GAC G IDLKTAKARLVTIACCI GA S A HT A HARHLVDHLIEEFGE---DFP P L Y I PF CD-----LCC -LC TY GK CD LS-----GGKR GAC G IGMEAQQSRIVLLAACI GA A T HI S HGRHLVDHLIEKMGR---RTP P F Y A PF CD-----MCC -LC TY GK CD LL-----G-KK GAC G IDAEAQQARTVLLASCI GT A A HA G HGRHMLHHVIKEKGK---DFP P F Y A PF CD-----MCC -LC TY GK CE LL-----G-KK GAC G IDAATQQARTVLLACLI GT A A HA G HARHLVDHLIERLGE---DYK V I A FPI CD-----MCC -MC TY GR CN LA-----EGRR GAC G IDIRSNTARFTALKTCI GA A C HA A HARHLVEYILEKLG----DVE P V Y T PM CD-----QCC -YC TF GP CD LS-----GNKE GAC G INLEGHNAREFMLRVIT GA A A HS G HGRHLLHHLIGLYGK---DHP P V Y T PM CD-----QCC -YC TF GP CN LE-----GNRR GAC G LDMKGQAAREFFLRCIT GC A C HS A HGRHLLDHIISIFGE---DMP P V L T PM CD-----QCC -YC TY GP CD LS-----GNKR GAC G IDMAGQTGREFFLRVIT GT A C HA A HGRHLLDHVIEVFGE---DLP P V I T PM CD-----QCC -YC TY GP CD LS-----GNKR GAC G IDMLGHNGREFFLRVIT GT A C HA A HGRHLLDHLIETFGE---DLP P F Y API CD-----MCC -LC TY GK CD LT-----GDRK GAC G LDIATQQARMVVIACCI GT A A HA G HANHVLHELIKWRGR---DHP L A V VPG--------------------------------DSEEERREV ARTIVTLCCE GT AGLIST ARLVV-ELLRQTGE---N--

Met. Ions Life Sci. 2, 357–416 (2007)


397

Appendix 2. (Continued) R . r u b ru m M o . t h er m o 2 C . p h a eo b C . d i f fi c i l 2 M . j a n n2 M . k a n dl 3 D . h a f ni e n 5 C . h y d ro 5 M s . a c et 4 S . f u m ar o x 2 C . h y d ro 2 D . h a f ni e n 2 A . m e t al l 1 C . h y d ro 1 C . h y d ro 3 D . h a f ni e n 3 D . h a l fn i e n 1 C . h y d ro 4 A . m e t al l 2 C . d i f fi c i l 1 G . s u l fu r M . h u t ch i n 1 S . f u m ar o x 1 M s . a c et 3 A . f u l gi d 3 M o . t h er m 1 M c . j a nn 1 ANME1 A . f u l gi d 2 A . f u l gi d 1 M . h u t ch i n 2 M c . m a ri p a l M . t h e rm a u t M . k a n d1 M . b u r to n i i M s . t h er m o M s . a c et i v 2 M s . a c et i v 5 ANME2 M . k a n d2

119 127 154 111 111 120 122 114 119 134 116 115 113 117 115 114 137 136 116 111 121 123 118 122 114 137 138 130 133 130 134 134 137 120 134 137 137 138 131 93

I R D E A K L Y A IAKTLGVATEGRGLLAIVGDLAAITLGDFQNQDYDKPCAWLAASLTPR RVKRLGDL------------GLLPHNID L K G E E R V L D VANRLGLTTDGRTIKEIAREVAEVLLEDLGRLTMT-PIRILTAYAPRE RIETWQTL------------GVLPRSGA I K C P E K G V A VARALGIETEGKTIEAICGEIADIFIDDLQRSLPK-RHETLHALAPKE RAELWERL------------GIIPISAY V K G E K T L N Q LAEMFGIESDEKYD--KCIKVADKVINDLYKSRDD-KMELVEKIAYAP RVKKWKEL------------GIMPGGAK I R N E K K L K F LAKKLGFDAN-KDAKQLAVEVAEFILDDMYKPRWE-KSELVPKLCPEK RLEVFEKL------------DILPGGAK I A D E K A L R HAAEVYGLDTSGKP-EDVAEEIAEFILEDIYRPRYE-ESEVFKAVVPDW RIEMYEEM------------GLIPGGAK I K E P E K L Q W MCEKLGIDTD-QDMNKMAIQLADLLEHQQQIGVEE-KNLMVEAFAPKK RKQVWRDL------------NIYPAGTV I T D K D K L Y Q MAKDLELNTEGKP-EDVAVRLSDFLIWELYRDYDE-PGKMIEVYAPLK RKEVWRKL------------GIYPAGPL I R E P E K L R TFAGRLGIETLGSD-SEIALYLCEFVEKDFNRPAYE-PSRIVEILAPPE RKKRWEEL------------DIFPGGIY I K N E K K L F E VAKSIDIKVDGRSVKDVATDLYTELERTYTQVEGE---IPFMKRVPPKTLELWRKA------------GIVPRGA M I K D E Q K L R K IAEKLNLAPAGKDIRQVAKEVALASLDDYSRQKRNVPCNWAKETLTAE RVDKLAEL------------GVMPHNID I K D E E K L K R IAQRLGIDWEGKSIRELTKEVADASLEDYSRQDSKIPCRWAELTMTQE RAAKLKQW------------DVMPHNID I K D S E K L R A IAKKMNISIEGKDIYQIAEQVAYESLKDYSRIEEE-PCTWLSNSITPP RLEKLEGL------------GVALNNIE I K D R T K L H S IAKRLGIPTEGQKDEDIALEVAKAALADFHEK-DT-PVLWVTTVLPPS RVKVLSAH------------GLIPAGID I K D Q E K L K Q VAFTLGIDTANKTEQEIVEEMCQIIYRDFANSGAT-PMTYLKANSPRE RLETWEKL------------GVLPRNPD I A D A G K L K A VASGLGIDSEGKEPDALALEVVRVAYGDMGNHHNR-PMKWVSAHAPKQ RLEVWESL------------GIIPRNPD I K S P E K L H K LAEKLGIETLNRDNMEILGDVTELAYEDFGRYKDR-PAAFLDSFLIEQ RHNKFLNT------------NIMPRNID I T D P D K L R R IAQRLGLITQGKDDMTLAKEVAELALEDFARLPGFGENLWIKTTLNKE RLEKYDEC------------NIMPSGIF V Q D E K K L I A LAQEWEVPTEGRDIYDVAHEVAEIALNEFGKPF---GTLRFPDRAPAP RRKIWEEL------------NIMPRAID I R D E A K L I T LAKEWDVETEGRDIYDVAHEVAEIALMEFGKPF---GTLRFIKNAPEP RQKIWKEY------------AIEPRAID I A D P D K L T A VAERLGIPTAGRSTAEIAGDVAAVAIDCFGNQGEE-PIVFMEKYMPKK RFQRLRELEETLYRTTGAKTGLLPRAID I K D V E K L H R LATEYGIQTDGRSPNDIAKDLALGILEEFGTIK---NRIQLAELAPDKTKDIWRKT------------GIMPRSID I R D A E K L K R VASEYGISSDGKDDLKIAEQLAYAMQEDFGTRK---KALTLLRRAPAK RRQVWDKL------------GIAPRGID I K D E A K L Y S LASEYGISAEEKSREEVAVELASALLSEFGKQE---GPILCTKRAPES RLKLWSEL------------GIEPRGID I K D V E K L K WFAKLLGIE--GEDVNELAAKVADFVISDLSSLE---KSRLVEIFAPEK RKELWEKL------------GIFPSGVF V K D E A K L K E VCRRVGIEVEGKSVLELAQEVGEKALEDFRRLKGEGEATWLMTTINEG RKEKFRTH------------NVVPFGIH I D L G - N E I E VE------------------------------------APIARTVTGI KPKTLGDL------------EKILDYCE I D F G - P E V D IE------------------------------------APLTRLVCGI KPKTIGDY------------DKVFRYIE V D L G - P E I DEV------------------------------------APLTELITGI KPKTLEDL------------ERALRYAE I D L G - G D V N VE------------------------------------APIIRTVVGI KPKTLGDL------------REALNWAE I D V G G S N V D VE------------------------------------APMIRLVCGI KPKTLGDL------------ETVLDYLE I D M G - K N I D IE------------------------------------APIIRTIIGK KPVTVSDL------------DESLSYVE I D L G - S N V D IE------------------------------------APITRTVMGK RPATLGDL------------REVMDYAE I D L G - S E V D VM------------------------------------TPIFETLVGF KPKTVSDL------------EKGLEYIE L D V G - A - T N II------------------------------------APNVQLVTGVQPKTLGDL------------DSVLSYVE I N M G - A - S N VI------------------------------------APNIQLITGRQPKTLGDL------------KPIMEYVE L N L G - E - S N VL------------------------------------TPNVTICTGLSPKTLGEC------------RAPMEYVE L N L G - Q - S N VL------------------------------------TPNITISTGLSPKNLGEI------------KPAMEFVE I D L G - P Y I D VE------------------------------------APIIRTVVGT RPKTLGDL------------EEILNYVN L D P G - F D A ETP------------------------------------LPLYETLLGSSPECADDL------------EAGLSYAE

R . r u b ru m M o . t h er m o 2 C . p h a eo b C . d i f fi c i l 2 M . j a n n2 M . k a n dl 3 D . h a f ni e n 5 C . h y d ro 5 M s . a c et 4 S . f u m ar o x 2 C . h y d ro 2 D . h a f ni e n 2 A . m e t al l 1 C . h y d ro 1 C . h y d ro 3 D . h a f ni e n 3 D . h a l fn i e n 1 C . h y d ro 4 A . m e t al l 2 C . d i f fi c i l 1 G . s u l fu r M . h u t ch i n 1 S . f u m ar o x 1 M s . a c et 3 A . f u l gi d 3 M o . t h er m 1 M c . j a nn 1 ANME1 A . f u l gi d 2 A . f u l gi d 1 M . h u t ch i n 2 M c . m a ri p a l M . t h e rm a u t M . k a n d1 M . b u r to n i i M s . t h er m o M s . a c et i v 2 M s . a c et i v 5 ANME2 M . k a n d2

192 199 226 181 182 191 193 185 190 204 189 188 185 188 187 186 209 209 186 181 205 193 188 192 182 210 174 166 169 166 171 170 173 156 169 172 172 173 167 129

A SV A Q T M S RTHV GC DADPTNLILGG LRVA MAD-LD GSMLATELSDAL F--------G TPQPVVSAANLG VMKRGA-VNIAVN GHN Y E I M E T L H MTTL GGTSDWTSLTEQE LRAA LAYCYSTLFGSSLATEML F--------G IPRPKVATVNYG ILKEDH-VNILIH GHS H E C F E V N N LTSHGT DSDFESHMQAF LRTV LAYAITTVTSTSLATDIV Y--------G LPRRSKLNVNLG SIVPDGCVNIGIN GHA S EV F D A I V KSSTNLNSDPVDMLVNC LNLG ISTGLY GLTLTNLLNDVML-------- G EPVIRMAPVGFNV IDPDY-INIMIT GHQ G EI V D A L T KTSTNLNSNPMDLLVHC LRLG LHAGFT GLLMTCWLNDIL F--------G SPKITVVENGFSS VKPNN-VNIMIT GHQ S EI H D A L V KTSTNLNSDPVDMLLHV LRLG LITGPV ALFGVETINDIL F--------G SPKITQTEGGPG ILDPDY-VNIMTT GHQ H E E Q N C V A SCLTNVDGNYASLALKA LRLG LATIYNSQIGLEMVQDIL F--------G TPQPHEVDVDLG IMCPEH-VNIVFN GHQ H EL K D A A A SCLTNV DGDYVSLATKG LRLG LSCIYG AQIGLELVQDIL F--------G TGMPHEMDVDLG IFDADY-INIVFN GHE G EM M L S T S SCLTNV DGYYASLALKA MRLG IAMAYQSQIVNEYCQDIL F--------G IPKPHTMRVDLG VLDPEY-VNVLPN GHE R E I M E I M H RSHM GV DQDYENIVKQCSRTA LADGWG GSMVATELGDIM F--------G SPEPKKAGVNMG FLKEDQ-VNIIVH GHE A VI T E I M G RTHV GC DADAVNLLLGG IKGA LAD-YT GMCLSTELSDVI F--------G TPKPVITQANLG VLKEDA-VNIAVH GHN A A V A E I M S RTHV GC DADPVNILLGG VKGA VAD-YT GMYLSTELSDAL F--------G TPAPTVTEANLG VIKEDA-INIAVH GHN S S V A D I M S RTHL GT DADPINLLLAG MRCA IGD-YT GMQISTNLSDVL F--------G TPEPVISEANLG VIKKDA-VNIAVH GHN H EI A E I M H RTSM GC DADAQNLLLGG LRCS LAD-LA GCYMGTDLADIL F--------G TPAPVVTESNLG VLKADA-VNVAVH GHN R EI R E A L H QTTM GM DADPVNLILKT IRLG LVDGFA GLKLATDLQDII F--------G TPQPVVTEANLG VLKEDY-VNIIVH GHV R E I R E A M H QTTM GM DADPVNLLLATAKQG LVDGYA GLKLATDMQDIL F--------G TPAPVVTEANLG VLKEDY-VNLIVH GHV G TV T E L M A QTAQ GV DADPVNIIFGG LKGS LAD-LV GEYIGTNLSDVL F--------G IPEPIVSEANLG VIEEKM-VNIAVH GHN G DI S D L L A QAHI GN DDDPVNITFSA LRVA LTD-YA GMHIATDFSDVL F--------G TPKPIVTEANLG VLDANK-VNIAVH GHN R E I A T I M H STHM GCTSDAESLLRLS MRTS LANGWG GSMLGTELSDIV F--------G TSTPRMTEANLG VLEEKQ-VNILLH GHD R EI A T I M H STHI GCTGDIDSLIHMS LRTS MADGWG GSMIGTRLSDIL F--------G TPVPRRTEANLA VLEENK-VNIILH GHE R E A V D IL H RTHF GC DHDPLSLVAQS VRCS LSDGWG GSLIATELQDILL-------- G SPIIRPVKANLG VLEAES-VNVVVH GHE R E I V E A M H RIHM GV DADYVNLLLHA MRTS LSDGWG GSMAATECSDIL F--------G TPTPLTSITNLA TLSRDK-VNIVLH GHN R E T A E M M H RTHM GV DNNWQSLLLQA LRNA LSDGWG GSMIATELSDIL F--------G TPKPATTAVNVG VLKKDQ-VNIIVH GHN R EI V E C M H RTHI GV DNDATHILLHG LRTS LSDGWG GSMIATEIQDVL F--------G TPEPKKSTVNLG VLSHDK-VNVIVH GHE Q E L L T M G S SAMTNV DSNYVSLAKKS MSMS IATCMA AQIALETIQDIL F--------G TPMPHESHSDLG ILDPEY-VNIAVN GHE A S I S E L V N QAHM GM DNDPVNLVFSA IRVA LAD-YT GEHIATDFSDIL F--------G TPQPVVSEANMG VLDPDQ-VNFVLHGHN E QI T H L L S AAHT GQ EGDYLDFESKA LHAG MIDD-L AREAGDLAQIVA YNMPK----G DEDAPLIELGFG CIDKSK-PVILCI GHN E Q I V Q L A D AIHT GQ EGSYMDFESKA LHMG MLDS-LSKEAADIIQIAC YGMPTSLPGG GPDVPLVDVGIG TIDTNK-PVIIVI GHN E Q I V Q V V D AVHT GQ EGSYLDYESKA LHLG MLDS-L GKEIADIAQICA FGYPK----G EDNQPLIEVGMG VMDRSK-AMILVI GHH K E I V K V L H STHI GN EESLLDYESKA MHVS MADH-V GMEVADIAQIVA YNFPKA----EPDTPLVDTGFG IVDKSK-PTIVVV GHN S Q V T E L L A AAHT GQ ESDPLDFESKVFHAG MLDH-V GLELADMAQISA FNLPKA----DPDAALVELGLG TIDAEK-PVILVI GHN E Q L S H L L S ACHT GQ EGNSLDFESKA LHAG MMDD-L AREVADITQIVA YDMPR----G DGDEPLVELGLG TVDKDK-PVVLCI GHN E Q M S H L L S ACHT GQ EGDSKDFESKAFHAG LMDD-LTREVADLAQIVALDLPK---- G DEDAPLVELGFG TIDTEK-PVVLCI GHN R EL T K V L S SVHV GQ EMDPHDYESKA LHAG MIDN-L ALEIADVAQIAA FDMPK----G E--APLVEFGPFAADDSK-PCILLV GHN E Q I T Q L L A AIHV GQ EGAAIDFESKA LHGG MIDH-V GMEISDIAQISCLDFPKS----DEEAPLADIGMG CLDASK-PTLIVI GHN E E L G Q L L A TVHA GQ EGAAIDYDNKA MLAG ILDH-V GMEVSDIAQVTALGFPKS----DPEAPLVEVGMG TLDASK-PVIIAI GHN E Q L T Q L L A TIHA GQ ESAEIDYDSKA LFSGSLDH-V GMEVSDIAQVSA YDFPKA----DPEAPLIEIGMG SIDKSK-PLIVAI GHN E Q L T Q L L A TVHA GQ ESAEIDYDSKA LFSGSLDH-V GMEISDVVQVAA YDFPKA----DPEAPLIEIGMG TIDKSK-PFLCVI GHN Q Q L V H V L S STHT GQ EGSYIDFESKA LHVS MIDN-L AKEVADIAQIAG FDFPK----G DPDAALVDFGFG AVDRTK-PIIVFI GHY R EL T S S V S ELLRSH--SLKGYESVA MHAGAIGL-L AMEIADATPSTLMEVTES------EE-VFEIGTDD LPR-R-PTVLLV GHL

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398

LINDAHL and GRAHAM

Appendix 2. (Continued) R .r u b r u m M o. t h e r m o 2 C .p h a e o b C .d i f f i c i l 2 M .j a n n 2 M .k a n d l 3 D .h a f n i e n 5 C .h y d r o 5 M s. a c e t 4 S .f u m a r o x 2 C .h y d r o 2 D .h a f n i e n 2 A .m e t a l l 1 C .h y d r o 1 C .h y d r o 3 D .h a f n i e n 3 D .h a l f n i e n 1 C .h y d r o 4 A .m e t a l l 2 C .d i f f i c i l 1 G .s u l f u r M .h u t c h i n 1 S .f u m a r o x 1 M s. a c e t 3 A .f u l g i d 3 M o. t h e r m 1 M c. j a n n 1 A NM E 1 A .f u l g i d 2 A .f u l g i d 1 M .h u t c h i n 2 M c. m a r i p a l M .t h e r m a u t M .k a n d 1 M .b u r t o n i i M s. t h e r m o M s. a c e t i v 2 M s. a c e t i v 5 A NM E 2 M .k a n d 2

267 275 303 257 258 267 269 261 266 280 264 263 260 263 263 262 284 284 262 257 281 269 264 268 258 285 253 249 248 245 250 249 252 233 248 251 251 252 246 202

P M L S DIICDVAAD--LRDEAIAAGAAEG I NI IG IC C TGH E VMMRH-G-VP-----LATNYLSQELPIL T GALE A MVV D VQC IMPS P V M V EK ILEKIRTPEIQELARKAGAK-G I VV GG MC C TGE E LLARY-G-VP-----TVTNIMGQELALG T GAVD T VVV D MQC VIPG P M V A FAICDIVGTPKIMEKVKRAGAD-T I RL YG MC C TGG EFIERDLN-IP-----LVAMASSAEMAVA T GAFD A IVV D QQD VLPG H S T F ANFQDRLKDEDVIKLAQSVGAK-GFK L VG CT C VGQ D LQLRG-EHYQEVFAGHAGNNFTSEAVLS T GAID I VLS E FNC TIPG H A L I QP LCEAAMEEDLIKMAKEAGAD-E I KI IG AT C NGQ D METRI-AHLPESFVGYIANNFTTEPLVA T GLID A VVS E FNC TFHG M A L M KYLTDAAEK--LEEEAKAAGAK-G I RI IG AT C VGD DFEARA-EHLPDTYAGFAGNNFATEALAAT GLVD A IVS E FNC TFPG P W I G AA MIERARSSDVQEKARAAGAK-G L RV VG SIE TGQ E LLQRF-E-MDDVFVGLMGNWLAIEPLLA T GTVD V LAM E ENC SPPA P F V G VALILAAKEAVNQDKAKAAGAK-SL RI YG SIE SGQ E VVQRF-Q-KDEVFRGLTGNWLTIEPMLA T GAVD V LAM D MNC SPPN P F L G FAMVQLARKPEWQEKAKAAGAK-GL RV IA S I ETGQ E MIQRW-E-EDDAFYGFTGNWISQEAVLA S GSVD LFAA D MNC SLPV P L L F EA MIVASSDPENLKKAEAAGAK-GI NL LG MC C SGA E VLGRH-G-VP-----HAGNFLSTEPVLA T GAVD A MAV D VQC IMTG P L L S EIICDVALK--MNEEAKKAGAKEG I NV VG IC C TGN E VMMRR-G-IP-----LATNYLSQEMAII T GALD A MVV D VQC IMPA P L L S EV VCDAAAM--LNDLAKQAGAPGGFN I VG VC C TGN E VMVRH-G-VP-----LATNYLSQEMPIL T GALE A MVV D VQC IMPS P L L S EI IVDVVDE--MQQEARAAGATAG I NI VG VC C TGN E VLLRR-G-IP-----MASNYLSQELTIL T GALE A MVV D VQC IMPS P V L S DIIVSVSKE--MENEARAAGAT-G I NV VG IC C TGN E VLMRH-G-IP-----ACTHSVSQEMAMI T GALD A MIL D YQC IQPS P L L S EKIVEWSRK--LEDEAKKAGAK-G I NL AG IC C TGN E VLMRQ-G-VP-----LATNFLAQELAII T GAVD L MVV D VQC IMPS P L L S EK IVQWAKT--LSPEAEAVGAK-G V QV AG IC C TGN E VLMRQ-G-VP-----LATNYLAQELAIV T GAVD A MVV D VQC IMPS P V L S EMVVGAARE--LKAEAQKAGAE-G V NI VG IC C TGN E LLMRE-G-VY-----LATSSASQEMAIL T GVLD A MVV D IQC IYPS P L L S EKVVDAAKE--LEEEAKAAGAE-G I NI VG MC C TGN E VLMRR-G-VH-----LATSFASSELAIV T GAMD A VVV D VQC IMPG P C L S EM IVAAAQDPEMIKLTEEVGAE-G I NL AG MC C TGN E VTMRH-G-VK-----IAGTFYQQELAVL T GAIE A VIV D VQC IFPS P A L S EMIVLASEEPDLVALAKEVGAD-G I NL AG MC C TGN E ITMRH-G-VK-----IAGDFHQQELAII T GAVE A VIV D VQC IFPA P I L S AKVVEMAQSPECRAAAEAVGAK-RV NV VG LC C TGN E VLLRQ-G-VG-----MAGNESHSELAIM T GAVD A MVV D VQC IYPA P L L S QM IVNVAEEPDLKKMAEEKGAK-G I NL VG MC C TGN E LLMRS-G-IP-----LAGSFFDQELAIA T GAVE A MVV D YQC IYHS P V V S EM VLLACRSEDLLQLAASKGAA-G I NL AG VC C TGN E LLMRS-G-VA-----MAGNHLTTELVLT T GAVD M MIV D YQC IMPS P I L S EMIVEAAEDPELLELAEEKGAT-G I NV AG IC C TGN ETLMRH-G-TP-----IAGTFLQQELAVI T GAVE A MVV D VQC IMPS P F V G IA LIKLAEREEIQEKARKAGAK-G L RI IG FIE TGQ E ILQRV-D-SP-VFAGIVGNWIVQEYALA T GCVD VFAA D MNC TLPS P L L S EI IVQAARE--MEGEAKAAGAK-G I NL VG IC C TGN E VLMRQ-G-IP-----LVTSFASQELAIC T GAID A MCV D VQC IMPS V V P G SYILEYLEENSME---------DE V EV CG IC C TAI D ITRVSDK------PKVVGPLSRQLMFVR S GVAD V VIV D EQC IRTD V P P A AN IGVYLIENGLD---------DK V EL GG IC C TSI DTTRVYTK------AKVVSALGRQLRVLRA GIAD V VVT D EQC IRAD A P P V LN IADYIEENGLE---------DE V DL GG IC C TAN D MTRYYQK------AKIVSALGRQLKVIRA GLAD V IVI D EQC IRAD V M Y A RP VADYLEEMGRI---------DDFEL AG LC C TAH D MTRYNAK------AKIFGPISYQLRVIRA GIPD V MIS D EQC IRAD V P P A ID IIQYSRKHNLS---------GK I EV TG IC C TAI D LTRFDPN------AKIVGPISWQLRYIR S GVPD L IVV D EQC VRAD V A P G AE ILDYVEDNDLY---------ED V EI CG IC C TAI D ITRYNQA------AKVIGPLSKQMKFIR S GVSD V IVV D EQC VRTD V L P G AD IVDYLDENEME---------DQ V EV CG IC C AAI D VTRYNEA------AKVVGPLSKQLRFIR S GVAD V IVV D EQC VRTD V A P G TEVLDYLEERGLD---------EE V EV LG IC C TAW D VSRVDDR------SKVIGPLSRQLHYVRM GIAD V VVL D EQC IRAD V A A V TD IIDYMEDKGLN---------DK I EL GG LC C TAL D MTRYKTGDRTLPRAKVVGTLAKELKTIR S GIPD V IIV D EQC IRAD V A G V TY IMDYMEDNNLT---------DK M EI GG LC C TAF D MTRYKREDRKPPYAKIVGTISKELKVVR S GIPD V IVI D EQC VRAD V A G V TY IMDYMEENNLT---------DK M EI AG LC C TAF D MTRYKEADRRAPYAKIVGSLAKELKVIR S GMPD V IVV D EQC VRGD V G G V TY MMDYMEEHDLT---------DK M EI AG LC C TAI D LSRYKEADRRPPYAKVIGSMSKELKVIR S GMPD V IVV D EQC VRGD P A V S VATIDYIEEHG LS---------DK I EV CG IC C TAI ETSRYSAS------AKVIGPLSMQLFFVR S GIAD V IVL D EQC VRTD P L L G HIITE--ELGTLA---------RQ V EL VG LTH TAWP--NREDH------VRVVGPLSMYHEYLS S GFAD V VVV D GAC PGED

R .r u b r u m M o. t h e r m o 2 C .p h a e o b C .d i f f i c i l 2 M .j a n n 2 M .k a n d l 3 D .h a f n i e n 5 C .h y d r o 5 M s. a c e t 4 S .f u m a r o x 2 C .h y d r o 2 D .h a f n i e n 2 A .m e t a l l 1 C .h y d r o 1 C .h y d r o 3 D .h a f n i e n 3 D .h a l f n i e n 1 C .h y d r o 4 A .m e t a l l 2 C .d i f f i c i l 1 G .s u l f u r M .h u t c h i n 1 S .f u m a r o x 1 M s. a c e t 3 A .f u l g i d 3 M o. t h e r m 1 M c. j a n n 1 A NM E 1 A .f u l g i d 2 A .f u l g i d 1 M .h u t c h i n 2 M c. m a r i p a l M .t h e r m a u t M .k a n d 1 M .b u r t o n i i M s. t h e r m o M s. a c e t i v 2 M s. a c e t i v 5 A NM E 2 M .k a n d 2

343 352 381 340 341 348 351 343 348 357 340 339 336 338 338 337 359 359 339 334 358 346 341 345 339 360 323 319 318 315 320 319 322 303 324 327 327 328 316 268

L P R I AECFHTQ I IT TDKHNKISGATH---------VPF-----DEHKAV---ET A K T II R M AIAAFGR R DPNR----V------M K I V ADCFGTQ V IT TCNSNRIPGAIH---------IPF-----DPENPEGLDED ALKVARL AVEAFAH R DRSKM----------M M H V ARQFHTR V IT TSPSGRKEGAIV---------LELDYYLKNLDKIY---EL A E E IL D I AIDNYRN R ESKK----V------L E P I ADKYKVK M VCLDDVAKKANAEY---------IGL-----DRSKLD---ELSNTLI E K ALESYKE R RGSI-EIDIPK----L K F V AEKTKTK L ICIDDMAYVEGAEY---------IPW-----EPENAK---EK A R E II K K AIEAFKE R KGM----QK------L K F Y KEKLDVE L VAVDDVAKVWGAEL---------ILW-----DPERAE---EV A E E AV Q R AIEAFKE R RSKH-EDKI------I D M Y AEKYQAT L VSISTIIDIPGLQHK--------FPY-----DPSETD---KIVEALI E L AIDNFKK R KGKV-TP--------L G P L AEKYGAT L VSVSRLVRFPGIHHF--------LDY-----KPSEVR---EI A Q K II D I AVDSFKN KRHGKITP--------A P L Y AEKYGFKL MPVSELIAFEDITER--------LNY-----NPVEAG---RQ A A K LL N M AVENFKN R KNSG-EP--------L N T M SKCYGTRFFT TNRKAMIEGAEH---------VEF-----DEHDTE---KCTNQI I D M AIERFRN R P-VQ----V------L T S V AECFHTE I IT TMAENKITGATH---------IEF-----REDSAV---ES A K K IV E V AIEAFKK R DKRK----V------L G A I AQCFHTQ L IT TMSTTKIPGATH---------VQF-----DKENAV---ES A R K IL E L AVDAYQR R DPKR----V------L A K I ASCYHTK LFTTMPEAKIPGADH---------FEL-----HDENAV---EN A R K IV K Q SIEAYKR R DPNK----I------V A T I AECTGTT V IT TMEMSKITGATH---------VNF-----AEEAAV---EN A K Q IL R L AIDTFKR R KGKP----V------L A E I AACYHTR L VT TMPIVKIPGAEH---------VPF-----TTETAD---EASQQI V R M AIESYQK R NPAK----V------L S Q I SSCYHTE L IT TMPIVKIPGATH---------VPF-----ALEKAD---EA A Q E IV R K AIGAYAR R DPNK----V------L T Q L CDCYHTK M IT TEAIMKVPGAQH---------LAF-----NSETAM---ED A K K LV R I AIEAYKR R DPKK----I------L K Q V TECYHTR L IT TSNIAKMPGTYH---------VPF-----HIENAL---ES A K E IV R L GIEAFKQ R VGKP----V------L A P I TDCYHTKF IT TSPKAKITGATH---------IEF-----NEEKAL---ES A K V IV R E AIENYKN R KQDK----V------L A R V ADCYHTKF VT TSPKAKITGSTY---------IEF-----REEQAL---DD A K A IV K E AILNFKN R DKSK----V------L A D L ASCFHTKF VT TSEQAKIPGALH---------IQF-----EEHEAD---AI A T R II K T AIDAFPN R NKAR----V------I P L T ASCYHTK V IT TSTKGKISGTIY---------KEF-----RPENAR---DT A H E II K L AIENFPN R NPDR----V------L G T V AACYHTR M VS TSDKARFPGMEH---------REF-----HPDNAA---EE A R A LV K E AIENFTN R G--E----V------L G N L TGCYHTKF IS TSPKADFPGTAR---------MEF-----HEEEAY---AT A K E IV K A AVENFPN R NLKK----V------L P E Y Q-RYGVK I VPVSRLVRLKGIDEG--------LDY-----EPEKAE---EI A M K LI D M AIENFKQ R DKSKA----------I S A V AECYHTR I IT TADNAKIPGAYH---------IDY-----QTATAI---ES A K T AI R M AIEAFKE R KESN-RP-V------I L E E VLKTGAV L IA TNEKMCLGLEDVSHMDEDEIIGYLLRNR--AALLLDEKKV G K V AV E V AKIVAKE R ---KDRKTLPDLNEVV V L E L CQRVQSP M IA TNDKAMHGLVDRTDDDPDKIVDDLVSGRVPGVIILEPEQV G E V AV R T ALQIDKK R ---KGLNWILSDDEFT I L Y H TKKLGIP V IC TNEKAMHALPDMTKEEPKNIIKYLLDGN-PGCVILDPLKV G E V AV E V ARARRKQ R G--DDIGPRLTEEQFM L L E A CKKMGIP L IA TSDAAARGLPDVSDWPVEKIVDALVSGKLPGVFLPIPEKV GQVAPLV AEAIFKKHGGERKYKFFESDEALM L L I E AGNIQAP L IA TSSKNCAGLVDRTDDDPDDIVRDLVSGVVPGVLILNPLKV G E V AV R A ALESHEI R KT-KKTSIIPTAEELI V L E E AKRNGAV V IA TTDKMCLGLPDMTKEDPELIINELVNGNIDGALILEPKKV G K V AV E V AKKIAKR R ---ESLKCLPDISEIP V L E E ALKNRSA V IA TTDKMCLGLPDMTDEDPDKIVNDLINGNIEGALILDPEKV G E V AV K T AMKLAPI R ---KSLKKLPDIDEII I V E E ANEVGSR V IA TRDLVMAGLPDVTDEPTEKIIEKMVSGEWMGVFIEDLEKA A E V AV E V AIRVHER R K--K-EIPQPDPKKLQ V L K E ASKLMIP V IT TNDKVMYGLKDRSNDSIEDILEDLTTGKEKGALMFDYVKL GELAPRLTMMMSEI R KQ-KGIKALPTDEELK L V E E GKKLKIP V IA SNEKVMYGLPDRTNDDVDAIIEDIKTGKIPGCVMLDYEKL GELVPRL AMEMAPL R ---EGISAIPSDEEMA V L S E SQKLKIP V IA SNEKIMMGLPDRTDADVDSIVEEIKSGAIPGCVMLDYDKL GELIPKI AEVMAPI R DA-EGITAIPTDEEFK I V P E AQKLKIP V IA SNAKIMYGLPNRTDANVDDVVEELKSGAIPGCVMLDYDKL G E L CI R L TMEMGPI R DA-EGITAIPTDEEFA L L E Q VQKTGAP L IL TTDKICYNLPDVTNRDPDRVVSDLLNGA-PGVLISDPLIA AQVIAKT ALEIKPK R D---KLNHLPELDEVQ V I E A AREGGSK L VA TVGARVADLLDVTDYPVEEAVEVLVTEE-DAVYVEEPIKAVEIAAWA ALRVEGS R DR-R---------EPP

Met. Ions Life Sci. 2, 357–416 (2007)


399


400 412 443 402 398 408 410 403 407 413 397 396 393 395 395 394 416 416 396 391 415 403 396 402 396 419 403 401 400 400 404 401 404 385 408 409 411 412 397 342

- - - - - - - - ------------AIPAFKQKSIVGFSAEA VVAAL--------------------------------------A---- - - - - - - - ------------H IPRETTEAMVGWSYEA IVDTF--------------------------------------G---- - - - - - - - ------------HVPNIRAKVELGFSVEE VMKLF--------------------------------------N---- - - - - - - - ------------DHG--FEQSLTGVSEKN LKAFL--------------------------------------G---- - - - - - - - ------------DYYDEKVKSVVGVGEES LVEFL--------------------------------------G---- - - - - - - - ------------MEPKHRHENVVGFGYFS IEEAV--------------------------------------G---- - - - - - - - ------------K VPQYKAKAIAGFSTEA VLGAL--------------------------------------G---- - - - - - - - ------------KIPANIQKAITGFTPEA ILKAL--------------------------------------G---- - - - - - - - ------------VLNLPVKEAVVGFSTES ILDAL--------------------------------------G---- - - - - - - - ------------E IPQRQDWGIHGFSMEY ISYML--------------------------------------G---- - - - - - - - ------------NIPDCKQTAITGFSAEA IMAVL--------------------------------------S---- - - - - - - - ------------N IPQVKEKAIVGFSAEA VIGAL--------------------------------------S---- - - - - - - - ------------L IPEEKSQIIAGFSTES ITNLL--------------------------------------S---- - - - - - - - ------------EIPNIKTKVVAGFSTEA IINAL--------------------------------------S---- - - - - - - - ------------YIPREKAKVVAGFSVEA IVKAL--------------------------------------A---- - - - - - - - ------------H IPDYRAKIIAGFSVEA IAGAL--------------------------------------G---- - - - - - - - ------------A LSKTRNSLVAGFSIEA LTDIF--------------------------------------G---- - - - - - - - ------------HIPEVKHKVVAGFSFEA LMEIF--------------------------------------A---- - - - - - - - ------------F IPQSKAQAITGYSNEA VIKQL--------------------------------------DRVVN - - - - - - - - ------------L IPELKSGATVGYSVEA VVNQL--------------------------------------DRVVN - - - - - - - - ------------YIPQHTSTAIVGFTVEE ILKAL--------------------------------------G---- - - - - - - - ------------R IPERPMKVMAGFSEET IRKAL--------------------------------------G---- - - - - - - - ------------Y IPVEPVNAVGGFSVET ILGAL--------------------------------------G---- - - - - - - - ------------SIPEEKQECMVGFSAEA ILKAL--------------------------------------G---- - - - - - - - ------------VKVEQKKKIVVGFSPEA ILKAL--------------------------------------N---- - - - - - - - ------------Y IPQIKNRVVAGWSLEA LTKLL--------------------------------------A---E L A K Q C T E CGWCNRNCPNAFKVKEAMALAK-QGNFKGFIDLY--KRCYGCGRCEAICPRNLPIVSMTTKVGEAYYKDLKFKMRAG R Y V N D C N L CGSCTLACPTGLR IGEANKSAA-AGNVKP LADLF--DLCVGCGRCEQVCKKHIPIVDVVIKAAYPQLKEERGRMRVG E Y A R A C T Q CGNCTIACPQGIR IGEAMEAAE-NGDRSK LEKEW--DVCIACGRCEQVCPKGIPIIDMYNYAAWNLIVNEKGKLRRG E E I N K C T Q CMNCVFTCPHSLR VDQGMAHAQKTGDLSK LAQLE--EQCLACMKCEQACPKNIKIINVIMRANYDRLYNKTGKTRVG E Y A K R C G G CMECTRACPNETP LPEAMKQAA-TGDLSF LADIY--DTCIGCGRCDDACNKEIPIHSSLVAAAREKVLNEKHFVRAG E I A K E C T E CGWCVRVCPNNRP IMDAVTSAA-KGDLTK LANLYEHDMCYTCGRCEEECERNLPLVSFITKAGDHFVKNQKFKMRAG E L A S E C T D CGWCQRVCPNSLP VMDAVKKAA-DGDLSK LEEMAIEELCYTCGRCEQECERNIPIVSMVTKAGERRVKDEKYRIRAG K E A K R C L G CGDCERVCPNDLP IVEAMERAA-NGDFEG LADLF--DRCVGCARCESECPTKLRVMNMIEDAWRLRTKEEKYKVRTG E L A D S C V H CLKCEVACPNSLP ISEAMTALS-EGDLSKFELLH--DKCIACGRCEYACPKDIDIVNVIEKSSQRVISEEVGKVRVG S L V A K C V A CGECALACPEELD IPDAIQAAK-EGDFTA LDFLH--DLCVGCRRCEQVCNKEIPILSVIDKAAQKAIAEEKGLVRAG V Y I D K C V K CGECMLACPEELD IPEALEYAA-KGSYEY LEALH--DVCIGCRRCEQVCKKEIPILNVLEKAAQKSISEEKGWVRSG D W V A K C A D CGACMIACPEELD IPEAMGFAK-EGDFSY LEELH--DQCIGCRRCEQVCKKEIPILNIIEKVAQKQIAEEKGWMRAG K L A E E C I K CGWCDRACPNSLP VKDAMIKAK-EGDLSE LSNLF--SPCMSCGRCESECKKDLPIVSMILKSAQEIAFGETYTMRAG R R A F R - - - -----------------V------GPPT-------------------------------------------------


423 435 466 423 421 431 433 426 430 436 420 419 416 418 418 417 439 439 423 418 438 426 419 425 419 442 485 483 482 483 486 485 488 467 490 491 493 494 479 352

- K V N A D D PLKPLVDNVVNG NI QG IVLFVG CNTTK-VQQ DSAYVD LAKSLAKRNVLV LAT G C AAGAFAKAGLMTSEATTQYAGEGL - - - - - - G - LKGLLELLREG KI KG IATVVG CNTPK-VPY EFNHVT IVRRLI E S D ILV TTT G C CSHALLNAGLCSPAAAS-QAGPGL - G S M P D K KIHGLAALLKAG KI RG IVNFGS CG NVRGAVF ERNQII IAKQLI K N D VLV TAH G C SGMGLLFAGLAHPDASV-LCGTGL - - - - - - D S WKP LINLIAEG KI KG VAAVVG CS NMTAGGH DVNTVE LTKELI K K D IIV LSA G C STGGLENVGLMSPGAEE-LAGENL - - - - - - G S VKPLIELIASG KI KG VVGVVG CS NLASGGH DNIIVT LTKELI K R D ILV LAG G C VNSPLKHAGLFDPASAE-LAGENL - - - - - - W - -ENVLKLIEEG TI RG VCAIMG CT NLSSGGHNVPAVE LAKEMI K R D VLV LGA G C VNGAFANAGLFNPEAAE-LAGDNL - - - - - - N K LDPLVEVIAAG KI KG VVALAN CSTLRNGPQ DWNTVN LTKQLI K K D ILV VAG G C GNHALEVAGLCNLDAIK-EAGSGL - - - - - - G SINPLIEVIKAG KI KG AVGLIN CTTLKNGPQ DYVTVN LAKELI K R D ILI LSG G C GNHALEVAGLCNLDAIN-LAGPGL - - - - - - G TLDPLLDAIKSG AI KG VVGMVS CTTLRDYGQ DVHSVA VVKELI K R N ILV LSL G C GNGAMQVAGLCSPETRE-FAGDSL - G T F R S - S YVP LNDNIING RI RG VAGVVG CT NPR-VKQ DYIHVE LVKELI K N N VLV VQT G C SQIALAKAGLMKPDAAV-LAGDGL - K L N A N D PLKPLIDNIING NI QG IALFAG CN NPK-AIH DNSFIT IAKELAKNNVLM LAT G C GAGAFAKNGLMTQEATEAYAGESL - Q L D A H D P LQPLLDNIVNG NI QG ICLFAG CNSTN-TLQ DRSFVE LAKGLAAHNVLL LAT G C GAGALAKHGLMTQEATLAYAGDGL - K I D S E K P LRPLIDNIVNG NI RG VVLFAG CN NVK-ITQ DYNFVH MAKKLL E N N ILI LAT G C GAGAFAKNSLMNSEATNKYAGESL - K L N A N D PLKPLIDNVVNG NI RG VCLFAG CN NVK-VPQ DQNFTT IARKLL K Q N VLV VAT G C GAGALMRHGFMDPANVDELCGDGL - K L N P D D PLKPLIDNIVSG NI LG VVATVG CN NVK-VKH DWFHIE LVKELI K N N VLV VTT G C SAHALAKAGLMDPAAAE-WAGEGL - K L D P V Q P LKPLADNIVNG NI VG VVATVG CN NVK-VTQ DIFHVE MVKELL K N N VLV VAT G C SAHALAKAGMMNSEGTERYAGNTL - K I N P E R P FSV LTDAILSG QI KG VVQMAG CN NLK-RQQ DESHVA ILKELVKNDVF V VAT G C SAGAFAKMGLMNSEAVEEYAGEGL - H V N Q E N PIRVLNDAILSG QL KG VVLFAG CN NLK-RPQ DESHIT ILKEMLKNDVF V VTT G C SAQAFAKHGFLRPEALE-LAGEGL S N I D P Q G T VKPLADVIKAG VL RG AAGIVG CN NIK-TKH EYSHIE IMKKLI A N D VIV VTT G C AAQAAAKAGLLSKDAIK-LAGKGL S H I D P A G T VKPLTDCLKSG VL RG AAGVVG CN NAK-GVSNEAHVT IMKELI K N D IIV VTT G C GASAAAKFGLMESDAAEKYAGKGL - - - - - - G T PQPLIDLIVTG TI KG VAGIVG CN NVK-VQQ DFFHRT LTEELI K R D ILV IGT G C WAIAAAKSGLMDLPARE-LAGPGL - - - - - - G T YKP LIDAIVAG TI KG CVGIVG CN NPR-IKQ DYGHIT LAKELI K R N ILV VET G C SAIACGKAGLLVPEAAD-LAGEGL - - - - - - G T PQP LIDALKSG TI RG AVGIVG CN NPR-IRQ DFGHVT LAKRLI E N D ILV VDT G C AAVATAKAGFKASDAAQ-MAGPGL - - - - - - G S PAPLIEAIAGG AI KG IGAVVG CN NVK-IQHNYGHVN LVKELI K N N VLV VTT G C NAIACAEAGLLVPEAAA-LAGDGL - - - - - - G D LNVLLDAIKKG DI KG VVALVS CTTLKNGPH DSSTVT IAKELI K R D ILV LSM G C GNAALQVAGLTSMEAVE-LAGEKL - T Q N A Q N P IRVLNQAILDG EL AG VALICG CN NLK-GFQ DNSHLT VMKELLKNNVF V VAT G C SAQAAGKLGLLDPANVETYCGDGL R G P I K D V EIRSVGAPIVFG DI PG VVALVG CS NHPNGEE EV--AM IAKEFL E R K YIV VAT G C AAMAIGMWKDKDGK----TLYEKY R G P V W D S E IREVGAPLVLG TI PG IIAPIG CG NYPNGTK DA--WL IVKEFAERNYIV TLT G C MAIDCALWKDEEGK----TLYEAH R G P I R D S E IRNVGAPIVLG TI PG IIAVIG CG NYPNGTR DA--YT IMDEFASRNYIV VTT G C MAFDAALYKDEEGQ----TVYEKY R G P I Q D T E IRKVGQPIVFG QI PG VIAAVG CI NFPDEMKSI--RE ILEEFL K R R YIV VTS G C HAMDIGMIKDEEGK----TLYEKY R G A I Q D I E IREVGGPIVLG EI PG VIAFVG CA NFPNGAS EV--AE MAREFAKRRYIA VAT G C SAMAIGMYRNEDGQ----TPYEEF R G P V Q D V E IRKVGGPIVLG DI PG VVAFVG CS NFPNGGI EI--AK MAEELL K R R YIV VAT G C SAMSIGEYIDEDGL----TLYEKY R G P A Q D V E IRRVGAPIVLG DI PG VVAFVG CS NYPEGGK DV--AL MAKEFL E R N YIV VTT G C GAMSIGEYRDEDGQ----TLYEKY R G P I K D V E IRQVGGPIVMG DI PG VVAFVA CP NYPDDVKQV--GK MVEELL E R N YIV LTS G C TAMALGMYTDEDGK----TLYEKY R G P I S D P E IREEGVNLVLG TTPG IVALVG CS NYPDGTK DL--FT IADEML R R S YIV VVS G C SAMDLGMYKGEDGL----TLYEKY R G Q V S D A E IRAEGLNLVMG TTPG VIAIIG CA NYPAGSK DV--YR IAEEFL N R N YIV AVS G C SAMDIGMYKDADGK----TLYERF R G Q A S D A E IRKEGLNLVMG TTPG IIAIIG CP NYPAGTK DV--YL IAEEFL K R N YLLAVS G C SAMDIGMFKDEDGK----TLYEKY R G Q V S D A E IRAEGLNLVMG TTPG IIAIIG CP NYAEGTK DV--YY IAEEFL K R N FIV VTT G C GAMDIGMFKDEDGK----TLYERF R G P V L D T E IRKVGAPLVLG EI PG IVALVG CS NFPDGEV EV--AK IAEEFARRNYIV VAT G C SAMAMAMYRCEDGQ----TLYEKY - - R L T D V V IRNVGVPVVAG NI PG IVVLVS CPEKSADVE EP--AK IAEVLL E R G YLV LVP G C LAVALGSYLDDDGK----TLYERY

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Appendix 2. (Continued) R .r u b r u m M o. t h e r m o 2 C .p h a e o b C .d i f f i c i l 2 M .j a n n 2 M .k a n d l 3 D .h a f n i e n 5 C .h y d r o 5 M s. a c e t 4 S .f u m a r o x 2 C .h y d r o 2 D .h a f n i e n 2 A .m e t a l l 1 C .h y d r o 1 C .h y d r o 3 D .h a f n i e n 3 D .h a l f n i e n 1 C .h y d r o 4 A .m e t a l l 2 C .d i f f i c i l 1 G .s u l f u r M .h u t c h i n 1 S .f u m a r o x 1 M s. a c e t 3 A .f u l g i d 3 M o. t h e r m 1 M c. j a n n 1 A NM E 1 A .f u l g i d 2 A .f u l g i d 1 M .h u t c h i n 2 M c. m a r i p a l M .t h e r m a u t M .k a n d 1 M .b u r t o n i i M s. t h e r m o M s. a c e t i v 2 M s. a c e t i v 5 A NM E 2 M .k a n d 2

506 511 549 501 499 507 511 504 508 517 503 502 499 501 500 500 522 521 506 502 515 503 496 502 497 525 564 562 561 562 565 564 567 546 569 570 572 573 558 429

K G V L SAIGTA-AGLGGPLPLV MHMG SC VDNSRAVALATA L ANK--LG-----------VDLSD LPLVA S APEC-MSE K AL A IGSW Q E V C RSRG---------IPPV LAVG GC VDNTRTLRLFID L AEE--AG-----------VAMPK MPFVFV GPEP-GNE K TVGQ GVT R E V V QAKD---------IPPV LHVG AC TDSTRASQIMAYT ANA--AA-----------QPNPA MPFAMV AADP-AAE K TMGARYA K E V C KTLG---------IPPV LNFG PC LAIGRLEIVATE L AAE--LG-----------IDLPQ LPLVLS APQW-LEEQAL A DGAF K E V C KSLG---------IPPV LNFG AC LSIARIEQVAVA I AEE--LG-----------VDIPD LPVAA S APQW-LEEQAL A DATY R Q V C EELG---------IPPV LHYG PC LAIGKIEHLVFE I AEI--LREKTGEE-----IDIPD VPAVA S APQW-LEEQAL A DASS Q E I C SALK---------IPPV LSFG TC TDTGRISMLVTA L ADH--LD-----------VDIPQ LPIAVT APEW-MEQ K AT I DGVF S E V C RNLN---------IPPV LSFG TC TDTGRISLVVTA L ANA--LN-----------VDTAD LPVAVT APMY-MEQ K AT I DALF K A V C EALG---------VPPV LSYG TC TDTGRLADFLGA ISAV--MG-----------VPIPDLPIAA A APEY-MEQ K AT I DAIF Q E V C ETVG---------MPPV ISLG SC VDNSRILTACAE MVRTGGLG-----------DKLSDLPVAG C APEY-MSE K AI C IGQY K A V L TALGKA-AGLNGPLPLV LHMG SC VDNSRAVNVAVA I ANK--LG-----------VDLDK LPLVA S APEF-MSE K AV A IGTW K A V L TAIGQA-NGLNGPLPLV LHMG SC VDNTRAVSVAVA I AQK--LG-----------VDLDR LPLVA S APEA-MSE K AV A IGTW K A V L TMLGKE-AGLDAPLPPI F H MG SC VDNSRAVDLAVN L ANE--LN-----------IDLYQ LPVVA S APEV-MSE K AI A IGTW K A V L TAIGEA-NGLGGPLPPV LHMG SC VDNSRSVALVAA L ANR--LG-----------VDMDR LPVVA S AAQA-MHE K AV A IGTW R A V L TAIGTA-NDLGGPLPPV LHMG SC VDNSRIGDLVIA V ANY--LK-----------VSPKD LPIAA S APEY-QHE K AL S IGTW K A V L RAIGEA-AGLGAPLPPV LHMG SC VDNSRIGDLVTA L ANY--LG-----------VDSAA LPVAA S APEP-QHE K AL S IGTW K S F I RQLEEANPQLSTKLPLV F H LG SC VDNSRGMDLVQA M AKE--LG-----------VDTPK VPFAA S APEA-MHE K AV A IGSY K S F I KMLEEK-AGLQGQLPPAFFMG SC VDNTRASDILVA M AKD--LG-----------VDTPK VPFVA S APEA-MSG K AV S IGTW Q A V C ELVD---------IPPV IHLG SC VDNTRIIRLVSA VSEH--LG-----------VDNSDLPVVG V APEW-MSE K AV A IGTY A T V C KLVG---------IPPV LHMG SC VDISRILDLVGAA ANY--LD-----------MDMCD LPVVG I APEW-MSE K AV A IGCY Q A V C GQLG---------IPPV LHMG SC VDCSRMLNLAGA L ADH--LQ-----------VDISD LPLVG S APEW-TTE K AV A IGTY A S V C RLLG---------IPPV LHMG SC VDCSRILVLAAN I AKE--LG-----------VGIGD LPIGG A APEW-YSQ K AI S IGTY R G I C SALG---------IPPV LHMG SC VDNVRILVLASA L ANA--LG-----------TDISD LPIAG A APEW-YSE K AV A IGAY K G V C EALG---------IPPV LHMG SC VDISRILVLASA V ANS--LG-----------VDISD LPAAG A APEW-MSE K AV S IGAY K A V C KALN---------IPPV LSFG TC TDTGRAAYLVRL I ADA--LG-----------VDVPQ LPVAVT APEY-MEQ K AT I DAVF K G F L KRLGEG-ANIEIGLPPV F H MG SC VDNSRAVDLLMA M AND--LG-----------VDTPK VPFVA S APEA-MSG K AA A IGTW P G E F RAGG------------L VNCG SC LSNCHITGAAIK I ANIFAKVPLRGNYAEVADYILNK VGAVG V AWGA-MSQ K AA A IATG H G R F DAGG------------V LNIG SC VSNAHIHDAAIK V ASIFGGRPLRGNYEEIADYILSR VGACG V AWGA-MSQ K AA A IATG H D R F DGGG------------V VQIG SC VANAHIHGAAIK V ARIFAKRNIRANYEEIADYILNR VGACG V AWGA-YSQ K AA S IATG P G N F DAGG------------L VNTG SC VANSHIAGAAIK I ANIFAMRPLRGNYAEIADYVLNR VGAVGFSWGP-YSH K AA S IATG H G R F DAGG------------I VNVG SC VSNAHISGAAIK I ASIFAKRNLRGNYEEIADYVYNR VGAVG V AWGA-MSQ K AA A IAAG E G I F DASG------------L A N IG SC VSNAHISGACIK I ANIFAKKPLSGNFEEVADYILNR VGACG V AWGA-FSQ K AA A IAAG G G Q F DAKG------------L VNMG SC VSNAHVSGAAIK I ANIFAQKPLEGNFEEIADYILNR VGACG V AWGA-YSQ K AA A IATG E D R F DAGC------------L VNTG SC VSNAHILGACIK I AAIFAKKPLKGNFKEIADYILNR IGACGVLWGT-MSQ K ALAISTG P S R F KSGG------------ L LNTG SC VSNAHITGAVIK V ASIFAQKNISGNYEEIADYTLNR VGAVG V AWGA-YSQ K AA S IGTG P G R F ERGN------------I LNTG SC VSNSHISGTCHK V AAIFAGRNLSGNLAEIADYTLNR VGAVG L AWGA-YSQ K AA A IGTG P G T F AGGG------------L LNTG SC VSNAHISGAAEK V AGIFAQRTLAGNLAEIADYTLNR VGACG L AWGA-YSQ K AA S IGTG P G G F ECGG------------L VNIG SC VSNAHITGAAEK V AAIFAQRTLEGNLAEISDYILNR VGACG L AWGA-FSQ K AS S IGTG P G S F DAGG------------V VNIG SC VANAHVIGAAIK V AAIFARLPLRGNFEEIADYILNK VGACG I AWGA-MSQ K AA S IATG P D T - ----------------L LNTG PC TSAAHLVGACIR V GVIFGKLPIRGEFVRVADYVLNRVGACVI AWGGEYSEHLVSAAYG

R .r u b r u m M o. t h e r m o 2 C .p h a e o b C .d i f f i c i l 2 M .j a n n 2 M .k a n d l 3 D .h a f n i e n 5 C .h y d r o 5 M s. a c e t 4 S .f u m a r o x 2 C .h y d r o 2 D .h a f n i e n 2 A .m e t a l l 1 C .h y d r o 1 C .h y d r o 3 D .h a f n i e n 3 D .h a l f n i e n 1 C .h y d r o 4 A .m e t a l l 2 C .d i f f i c i l 1 G .s u l f u r M .h u t c h i n 1 S .f u m a r o x 1 M s. a c e t 3 A .f u l g i d 3 M o. t h e r m 1 M c. j a n n 1 A NM E 1 A .f u l g i d 2 A .f u l g i d 1 M .h u t c h i n 2 M c. m a r i p a l M .t h e r m a u t M .k a n d 1 M .b u r t o n i i M s. t h e r m o M s. a c e t i v 2 M s. a c e t i v 5 A NM E 2 M .k a n d 2

576 573 611 563 561 575 573 566 570 581 573 572 569 571 570 570 593 591 568 564 577 565 558 564 559 595 636 634 633 634 637 636 639 618 641 642 644 645 630 497

A V T IGLPTH VGSVPPV-----------------IGSQI VTKLVT----ETAKDLV G GYFIVDTD PKSAGDKLYAAIQERRAGLGL F L A H GISNV IGFPGPIPVPLPRPVAGAAPDEYDRGSNP VADFFA---GDGLYEKV G ARIYTEPY PKLAAQTIRMLIRRQRLALGW F V L NGIETYSCVQDNT-----------------LASDRFIDYVS----NRLRTIV G AAMNWNPD PYRTSEDILCMLDEKRAALGW G L A LGLPLH LALPPFI-----------------TGGKL VTEVLT----EKLKDLT G GHVIVNPD PKSSANQLEEIIIDRRKNLGL A V D M GFTVH VSPVPFV-----------------TGSEL VTKVLT----EAVEGLT G GKLIPEPN PYKAADLLEQTIMEKRKKLGI A L A LGITLH VSPVPPV-----------------TGSEL VTKTLL----EDLPDLT G GELIVETDMKRAGEILAEKIEEKRKRLGI A V A Y GAYTH LSPTPFI-----------------TGAPQ LVKLLT----EDVEGLT G GKVAVGDD PVEVANAIEAHIVAKRKGLGL A L A YGLYTH VAPDPPV-----------------MGAPN LVKLLT----RDLPSIT G GRIAVGSD PVKVADDILAHINDRRAKLGI A L A LGLYTY VNPVPTV-----------------TGAPD LVKLLT----EDCREVT G GVLNVEKDAVKAVDGIEQHIMEKRKKLGI F V A S GVYTVFGVTFPS-----------------VEGTKFHKLLF----EGLEEQGFGKWGFATD PYDIAHLMIEHIDKKRAALGI A V T LGIPTH IGIVPQI-----------------MGSSV VVEFLT----EKAKDLL G GYFIVETN PELAAAKLVAVIKERRRGLGI A V A L GLPTH LGTVPQV-----------------LGSQV VTEVLT----EKIKDIT G GYFIVETD PEEAVKKLFAVIQEKRAGLNL A A G L GMPVH IGLMPPV-----------------GGSPL VAEVLT----SGLKDIVDGYFIVEKD PLEAASKIIHVIEEKRAALNV A V T IGLPTH IGVFPPI-----------------TGSLP VTQILT----SSVKDIT G GYFIVELD PQVAADKLLAAINERRAGLGL A V A MGIMTH LGVVPPV-----------------VGSSK VTRILT----QDAEALI G GKFYVETD PYKAAAGIIEHIKAKRALLNL A V A M GITTH LGVIPPV-----------------LGSKQ VTELLT----VGLTEVI G GKFYVETE PQKAALGLIEDIRKKRQALGL C V S M GIPTH VGTMPYL-----------------EGSDL IYGIAT----QIAHDVF G GNFIFEVDEKIAAQKILNALEYRSWKLKI F V T LGVPVH VGTMPPL-----------------EGSELFYSITT----QIASDVY G GYFMFEVD PVVAARKILNALEYRTWKLGV V V S S GIDVF LGITPPV-----------------TGSPEFVDLLT----NKIEDMT G AKFFIDEN PLTLADKILERIEEKRTKLEA V V A S GIDTY LGIMPPI-----------------AGSSRAVDILT----SELKDKV G ATFTVNTN PKELAATIIEDIEKKRVHFEA F V G SGIPVH LWPLPPI-----------------LGGPQ VTKILT----SDAKDVL G GWFFVEED PKATADRMEQIIMERRAALGI F V S S GVYTV LGIPPKI-----------------FGSSN VLNLLA----DGVKGITNSAFAVEPD PLKTADLLEAEINRKRKALNI F V A S GVYTV LGPMPPV-----------------TGSMN VVKLLT----EGLQDVV G ATFSVEPD PEKAALAIRRHIEEKRRGLGL V V S SGVFTV LGTIPPV-----------------LGSQA VTALLT----KGLDGVI G ASFAVEPD PFKAADLMLEHIEGKRKALGL A V A Y GLTTH VSPVPPI-----------------TGSEDAVKLFT----EDVEKLT G GKVVVEED PLKAAELLEKVIEEKRKALGI W V S L GVPTH VGTMPPV-----------------EGSDL IYSILT----QIASDVY G GYFIFEMD PQVAARKILDALEYRTWKLGV V N R WGIPVI LGPHGAK-----------------YRRLY LSNG---EKFKVKDKKT G EILEIEPA PEHLIVTAENVKECIC----F N R L GVPAV VGPHGVK-----------------YRRAF MGRPDLKEDWTLIDATD G SKVYVEPG PQDLLVACETVEEAMP----F N R L GIPAV VGPHGSK-----------------YRRAF LGRPYNDEDWMVYDART G EKVRIEPA PQDLLVAAETIEEAIP----F N R L GVPVV VGPHGTK-----------------YRRAY IGKPWKKDKWWVYDIKSRQKVFIEPA PDSLLVAVETKEEAIV----F W R L GIPVI VGPHGAK-----------------YRRML LGRKDHDEDWYVFDSRT G EKVQVGPV PEHLFVTAETKEEAMV----V N R W GIPAV LGPHSSK-----------------YRRLY LGRIDKEEMWDLNDLRT G ETVKGEPA PEHLLYAAETMEEALV----V N R W GIPVV LGPHGSK-----------------YRRLF LGRADDEEKWKLKDLRT G EVIDGEPA PEHLLYAAENREEATV----F T R W GIPIVYGPAGLK-----------------YQTLY IGDL--DGDWTVYDART G KECK-EYC PIHLKYAAEDWREALV----C S R L GIPVI LGPHGSK-----------------YRRAL IAKPYEEEKWKVYDARN G SEMQIPAA PDYLLTTAETVEEMMP----C N M Y GIPAV LGPHSGK-----------------YRRAL IAKTYDENKWKVYDSRN G SELDIPPS PEFLITTAETWQEACV----C N I Y GIPAV LGPHSSK-----------------YRRAL IAKNYDESKWKVYDGRD G SEMTIPPA PEFLLTTAETWQEAIP----C N I L GIPAV LGPHSSK-----------------YRRAL IAKTYEEDKWKVYDARN G QEMPIPPA PEFLLTTAETWQEAIP----V N R W GIPVI VGPHGSK-----------------YRRAY ISNKEL-HEWELYDKRA G EVVKGEPA PEHLLYPAETMEEAIV----V T R W GIPVV LGPDPEA-----------------GS--L LV----EKNPKVIDACS G EEVE-DPT PEHLRCVVSDWKEAAI-----

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401


640 655 675 627 625 639 637 630 634 645 637 636 633 635 634 634 657 655 632 628 641 629 622 628 623 659 696 697 696 697 700 699 702 678 704 705 707 708 692 553

- - - - - - - - ----------------------------------------------------------------------------K - - - - - - - ----------------------------------------------------------------------------P V R D Y T I G TKEEIEDKIPDTVEIGR-----------------------------------------------------------K D - - - - - - ---VECNA--------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------H - - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------K G - - - - - - ---ERERKL-------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------T S - - - - - - ---TLEEEK-------------------------------------------------------------------P R - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------D Q - - - - - - ----------------------------------------------------------------------------H K - - - - - - ---QTAEKFDTAIAQGW-----------------------------------------------------------H K - - - - - - ---QTAEKFETALCQNYGKEGSIAKLLILIKFLKGQ----LNRVKNPKGFLKRFMKEQLRPVM-----------RKF R L - - - - - - ---AKE--IDYGVIKP------------------------------------------------------------L V - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------P A - - - - - - ---QKV----------------------------------------------------------------------K - - - - - - - ----------------------------------------------------------------------------- - - - - - - - ----------------------------------------------------------------------------H K - - - - - - ---EVAERYETKLCQGY------------------------------------------------------------ - - - - - - - ---MIPKLCMRPNDTPKGRANKIYHYVDVYEKYFGRMPPDLEKFVRTEKDIPFMMKDKIMAYLEEK-----GWKPLE - - - - - - - - ---MMAKLCFRPSDTDRGRSIKLTHYIDLNLKYLNAYPPDWHLFVRAASDLPIATRNELLKKLEDEQSWKIDWKTKK - - - - - - - - ---LMAKLCFRPNDTTQGRSIKLTHYIDLSLKYLKRMPDDWHLFVRTEADLPLAKKEELLKELEDKHGWKIDWQKKK - - - - - - - - ---QLARLCIRPNDTNQGRQIKLTHYIELHQKYYGDLPDDWAVYVRSEADLPLKMRDQLLKVLEEQYGWKIDWDKKK - - - - - - - - ---LIAKLCMRPNDTSRGRSMKLTHYIDLYKRLYGIMPDDLHLFIRSPADIPITMKEELLPILQER-----GLKDRI - - - - - - - - ---STVKMCIRPNDTPKGRQLKLSNYIDLYKKYFGELPPDLELFVRNERDIPITHKKEVMDDLTSK-----NWTPRE - - - - - - - - ---MIAKLCIRPTDTPKGRQMKLSNYIDLHRKYLGTIPDDIDRFIRTEKDIPIVYKRDVMKILEEK-----NWKPRE - - - - - - - - ---QAVKLCIRPNDTPQGRQTKLQNYIELYKEFYNELPPDLPLYVRDKNDVPITLRDEVMEYLEEV-----GWKPRK - - - - - - - - ---MLAKSCIRPSDNNMGRMIKLTH---------------------------------------------------- - - - - - - - ---LLAKNCIRPSDNNMGRSIKLTHWIELSEKYLGVLPEDWWKFVRHEADLPLSRREELLKKLETEHGWEIDWKKKK - - - - - - - - ---MMAKACIRPSDNNMGRSIKLTHWMELSKKYLGVEPEDWWKFVRNEADLPLAKREELLKRLEAEHGWEIDWKRKK - - - - - - - - ---MMAKACIRPSDNSMGRSIKLTHWMELHKKYLGKDPEDWWKFVRNEADLPLAKREALLKELESKHGWEIDWKKKK - - - - - - - - ---AIAKFCIRPSDNAVPA------------RFIPTNPXG---------------------------G--------- - - - - - - - ---TAARLCMRPNDTPEGRQNKVESYVELYRELYGELPPDLDLLIRDESDIPVTLRSEIRELL-EETGWT----PRS


- - - - - - - - -------------------------- - - - - - - - -------------------------7 0 0 - - - - - - - - -----SICTII--------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------6 5 3 F S M E - - - - --DRRAAEG----------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------7 1 6 S F P G P L K N MDPTIRVILILPIFCRFGLLAVKKSE 6 4 5 - - - - - E E E VEETKELA-----------------6 3 0 - - - - - E E K MAEKAE-A------------------ - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------- - - - - - - - -------------------------7 6 5 K Y P Q - - - - -----DPTILY--------------7 7 1 I V E G P I R A YNPSFNPTNLERLIRRKR-------7 7 0 I V E G P I R G YHAGFNPTNLERCLRDGF-MT-V--7 7 1 I V E G P V R H FDAGFNPTIVEEVYEKYA-GEKAPR7 6 9 I P - - - - - - -----DPTLLKSQIRTQM-GGRS--7 6 8 A P K - - - - - -----EPSLLER----K--------7 7 1 L P K - - - - - -----EPSLLER-------------7 4 7 G I T - - - - - -----EPTLLEENVRG---------- - - - - - - - -------------------------7 7 9 I I S G P K I K FDVSSQPTNLKRLCKEA--------7 8 1 I I S G P K I K FDVSAQPTNLKRLCKEA--------7 8 2 I I S G P K I K FDVSAQPTNLKRLCKEA--------- - - - - - - - -------------------------6 2 2 R A S - D - - - ------PTLLPEG-------------

Met. Ions Life Sci. 2, 357–416 (2007)

1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

6 4 6 1 1 1 2 2 2 2 2 1 75 75 96 93 93 81 81 66

6 4 6 1 1 1 2 2 2 2 2 1 159 159 183 180 177 176 178 1 50

A NME3 A NME1 A .fulgidus M s.bark M s.thermo M s.acet M .burtonii M .hutchins M .kandleri M c.maripal M .thermaut M c.jann1 A .metallir C .difficil C .hydrogen M o.thermo D .hafniens D .ethenog S .fumaroxi M c.jann2



Met. Ions Life Sci. 2, 357–416 (2007)

- ---------------VTEEEEQAKDRQE-----------------------LWDSL--------------------------------- - - - - - - - - - - ------------------------------------------------------AE--------------------------------- - - - - - - - - - ----------------RI---EIPEGVKV-----------------------KESVG-------------------------------- - - - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - ---------------------------------------------------------------------------------------- - - - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - --------------------------------------------------------T-------------------------------- - - - - - - - - - - ----------------DK---LLRKNVKN-----------------------LREIL-------------------------------- - - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - ------------------------------------------------------------------------------------------ - - - - - - - - - GLLTFLVGE-----VIDQ---AIEANVKMGLELRVIPLGYDVTSVIHVVSVAIRASLIFGGVEPGNLEEHLAYTKKRVPAFVNAFGPLS E L V V S A G A G A I GLLTFLVGK-----VIDQ---AIEAGVKMGLELRVIPLGYDVTSVIHVVSVAVRAALIFGGLTPGDLNGLLEYTANRVPAFVNAFGPLS E L V V S A G A G A I GLMLFLCDE-----IIEQ---LLEENVKLGVDYIAYPLG-NFTQVVHAANYALRAGLMFGGIAPGLRDAHRDYQRRRVLAFVLYLGEHD M V K T A A A M G A I G FMLFICDE-----AVEQ---LLEENVKLGIDYIAYPLG-NFTQIVHAANYALRAGMMFGGVTPGAREEQRDYQRRRIRAFVLYLGEHD M V K T A A A F G A I GFMLFLCDE-----VIEQ---LLEENVKIGEDYIAFPLG-NFTQVIHAVNYAFRAGLAFGGIPAGQREQHRDYQHRRVRAFVLHLGELD D V K V A A E M G A I N LYTFMCGESQGKRFAKQ---LADADVELGWTPRLVSFGDEATAAIFAFGFAVRVALSFGNIEPGNRAKLFEYSQERIKAFIMPLGDIS D E W Y A N A A G A L NLYVFMCANQSGTTFSEQ---LIEAGVQIGWSTRLVPFGPDISAAVFALGFANRAAMAFGGVQPGDYATVLKYNKDRVFAFVNALGEVN A E W A A N A A G A I NILALLVGD-----IVKE---MDEADIEYGLDKLLVPVGNEITSAIHAANLAIRAPLIFGGIEPGKTEEIIDYLKNRVPAVVVALGELD N I T L A A G A G C I

- ---------------------------------------------------------------------------------------- - - - - - - - - - - - ---------------------------------------------------------------------------------------- - - - - - - - - - - ------------------------------------------------------------------------------------------ - - - - - - - - - - ---------------------------------------------------------------------------------------- - - - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - ---------------------------------------------------------------------------------------- - - - - - - - - - - ------------------------------------------------------------------------------------------ - - - - - - - - - ------------------------------------------------------------------------------------------ - - - - - - - - - ------------------------------------------------------------------------------------------ - - - - - - - - - ------------------------------------------------------------------------------------------ - - - - - - - - - ------------------------------------------------------------------------------------------ - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - - --EPRLEPALNAGLATAVSAEIIEAIKYALN---DAPY----------EAPCVGHISDATIRALGVPLVTGDIPGVAVVLGKCPDSETA G K V I K D Y Q S K - --THLLEHAFNAGLATALAAEVIEALKYSTM---DAPY----------SEPCAGHITDPIIRSLGVPLVTGDIPGVAVVLGECPDAESA A K V I K D Y Q S K - --ELTFENARLAGEATWYAAEIIEALRYLKHTPENPIV----------VPPWTGFIGDPVVRQYGIKMVDWTIPGEAIIIGRAKDSKAA K K I V D D L M G K ---VLNFENARLAGEATWYAAEIIEALRYLKYKPDEPLL----------PPPWTGFIGDPVVRRFGIKMVDWTIPGEAIILGRAKDSKAL A K I V K E L M GM - --ELTFENARLWGESVLYAAEIIEILHYLRN--DEPK-----------VAPWTGFLGDPIVRKHGIKMVDWTIPGVAVILGRAKDSPS A K K I V D N L M G K AEDVTELGAVLEAGLSTLFAEEMLEALRYLSQ---PEYYTNTEDPTP--DNIWLGAADDIILRKRGVEFVDGTAPGFAAIIGAAPDNAA A E S I A A E L Q Q K K IYVPYLGHTLDAGMAAILAEEVAEAIRYVED---PDFYLPSEDCDVERGRIWLGAADDTVLRKRGIEFVDGRAPGFAAIVGAAPNPQI A K E I A E D Y Q K K - --EETLENALDAGVVTLICAEAIEALKYAKS---EKPYK----------EPYVGFIPDEILRGLGVPLVEGKIPAILVVIGKVGDKEKL K K L I D D I K K R

M -------------------------------------------------------------------------------------TKE K - - - - - - - - - M -------------------------------------------------------------------------------------GE- - - - - - - - - - - M -------------------------------------------------------------------------------------VER K - - - - - - - - - - ---------------------------------------------------------------------------------------- - - - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - ---------------------------------------------------------------------------------------- - - - - - - - - - - M ---------------------------------------------------------------------------------------- - - - - - - - - - - M ---------------------------------------------------------------------------------------- - - - - - - - - - - M ---------------------------------------------------------------------------------------- - - - - - - - - - - M ---------------------------------------------------------------------------------------- - - - - - - - - - - M ---------------------------------------------------------------------------------------- - - - - - - - - - - ----------------------------------------------------------------------------------------- - - - - - - - - - - M ----------------------NLFQTVYNGSNVALNAAEGLLKQAIKEKGKDHNVAFPDTAYSLPIIYAATGLKMATLADLEGAIGV V K S L I V E - - - M ----------------------NLYNIIFTGSEQALGAAQAMLAEAIEKNGKEHKVAFPDTAYSLPCIYAATGQKMNTLGDLEGALEV V K S L I N R - - - M SEVINFDQIFEGAI-EPGKEPKRLFKEVYEGAITATSYAEILLSRAIEKYGPDHPVGYPDTAYFLPVIRAFSGEEVRTLKDMVPILNR M R A Q I K S - - - M ---TDFDKIFEGAI-PEGKEPVALFREVYHGAITATSYAEILLNQAIRTYGPDHPVGYPDTAYYLPVIRCFSGEEVKKLGDLPPILNR K R A Q V S P - - - M ----SMEQIYADAIKDPSKEPKKLLRKAYDGTITAMSYAEILLNRALKDYGPQQTVGYPDTAYHLPVITCLSGDKVTTLGELVPLLNR V R N Q V K T - - - M SKT--------------------VAEAAIKGARNIVNASYEVYKQTLVTNCPDTAVGFPNTAYFLPVIYGILGIQVEKLSDMGMVFDK C F K M L P P P E VN M SRL--------------------VAFAAIQGAYNIVSKAEGKFKTAMDKYGANQPLAFPNTAYYLPVIYSVLGIKVEKLADAEQVLNR C K S L L P P H I K D M -------------------------GNIIEGGKTVLNLT----KEILEKEDENLKVSYPGTNYNLPIIYGLLGKKIETVKDLKELINS L - - E I K D - - - -

402 LINDAHL and GRAHAM

Appendix 3. Alignment of α Subunit Protein Sequences

24 6 21 1 1 1 2 2 3 17 2 1 25 1 25 1 27 4 271 26 8 273 27 5 242

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181 150 16 4 142 142 142 14 4 14 5 14 5 15 7 14 1 140 43 6 43 7 46 1 458 45 5 463 46 6 422

A NM E3 ANME1 A.fulgidus Ms.bark Ms.thermo Ms.acet M.burtonii M.hutchins M.kandleri Mc.maripal M.thermaut Mc.jann1 A.metallir C.difficil C.hydrogen Mo.thermo D.hafniens D.ethenog S.fumaroxi Mc.jann2

ANME3 AN ME1 A .fulgidus M s.bark M s.thermo M s.acet M .burtonii M .hutchins M .kandleri M c.maripal M .thermaut M c.jann1 A .metallir C .difficil C .hydrogen M o.thermo D .hafniens D .ethenog S .fumaroxi M c.jann2

ANME3 ANME1 A.fulgidus Ms.bark Ms.thermo Ms.acet M.burtonii M.hutchins M.kandleri Mc.maripal M.thermaut Mc.jann1 A.metallir C.difficil C.hydrogen Mo.thermo D.hafniens D.ethenog S.fumaroxi Mc.jann2

-L V EKADAKIMIA TEDLGGSELTKGIIDNVCAPYWERV DK RA R A F SD DDA D TFY GCT I CQ TF APS H VC AVA PD RPPYC GII TWIGA KVMC DL D P YG Y I F E -F I ERMEA--TFI TDEEE VKK----RLE-EA NQIYEAR DM RSRG LHD EDV D EFY QCT L CQ SF APT N VC VVT PD RIALC GAM SWFDS RAAA RV D P EGP N A P -FI EKIEA--VYI TDKDL VEK----LLNELA MPIFEER DA RVEA LSD EDV D EFY SCT L CQ SF APT N VC IVS PD RPSLC GAI TWFDG RAAA KV D P EGP N R A -FI EKMQ V -- TF Y TDEAE VEK----QLE-EA KEIFKIR DA RTKD LHD EDV D VFY GCT L CQ AF APT N VC VVS PD RISLC GAI NWFDG RAAA KV D P EGP Q F A -FI EKMQ V -- TF Y TDQAE VEK----QMA-EA MEIFKAR DA RTKD LHD EDV D VFY GCT L CQ SF APT N VC VVS PD RVSLC GAI NWFDG RAAA KV D P EGP Q F A -FI EKMQ V -- TF Y TEQAE VEK----QME-TA KEIFKAR DE RTKD LHD EDV D VFY GCT L CQ SF APT N VC VVS PD RVSLC GAI NWFDG RAAA KV D P EGP Q F E -FI EKVEA--VYI TDLAE IEK----EMD-NVKAIYKSR DD RTRD LHD EDV D TFY GCT L CQ SF APS N VC VIT PD RISLC GAI NWFDG RAAA KV D P EGP Q F A -II EKIQ I -- TF F TGG EE FQS----TYN-LA RERYETR DA RARG LLD EEV D TFY GCA L CQ SF APS H IC VIT PQ RYANC GAI SWFDG RAAA RM D P DGP I F P NVI DSVEV -- TI M TDEEK VEE----FLE-YA RRVYKKR DE RAKG LSE EDV N EFY VCL M CQ SF APT H VC VIT PD RPSLC GSI TWHDA KAAY KI D P EGP I F P -II EAISI -- TI F TEEEK VKE----FVE-KA VSEYQAR DS KARD LSE EDV D VFY GCI M CQ SF APT H VC IIT PD RPALC GGI NWFDC RAAA KI D P EGP I F E -II ESIAV -- TL M TDEAA VQE----FLE-TA REKYETR DS RARE LSD EDV D VFY GCL M CQ SF APT H VC IVT PD RTALC GAI NWFDC RAAY KM D P DGP I F E -FI EKCD V -- TI I TDPEK VKE----ELE-KA REIYNKR DE KTKA LHE EDV D VFY GCV M CQ SF APT H VC VIT PD RPALC GGI NYLDA RAAA KI D P NGP I F E SVV DKCQ V -- TI I TDAAK IAE----MKKTDA EAIYMAR DE RLAS LTD ESV D TFY SCL L CQ SF APA H VC IVT PE RLGLC GAV SWLDA KATK EL D P AGP C Q P SVV DKCE I -- TI I TDAEK VSE----LKG-EA IAKYNAR DE RLAS LVD ESV D TFY SCN L CQ SF APA H VC VVT PE RLGLC GAV SWLDA KATK EL D P TGP C Q P SIV DRVQ V -- TI Y TDEQK VLE----LRE-IA RKKYAER DA RLRE LSD EAV D TYY SCL L CQ SF APT H VC IVS PE RVGLC GAI SWLDA KAAY EI N P NGP N Q P AIV DRVQ V -- TI F TDEAK VKE----YME-VA REKYKER DD RMRG LTD ETV D TFY SCV L CQ SF APN H VC IVT PE RVGLC GAV SWLDA KASY EI N H A GP N Q P AIV DRVQ V -- TI Y TDQQA VEE----NLV-KA RERYRAR DA RLKS LTD ENV E EFY SCL L CQ SF APN H VC IVS PE RVGLC GAV SWLDA KAAF EI T P TGP N Q P GIV DKIQ I -- KL Y TTP EK VAE----ISR-QSKKIYTKR DE RIEG MTD ENT E TFY SCT L CQ SF APS H VC IVS PE RTGLC GA Y NWMDC RASN EI N N I GP N Q P AIFDKCQ V -- KI Y TDEAK VNE----IIP-KA QAAYRVR DE RIEG MTD EGT E TYY SCT L CQ SF APT H VC VIS PE RTGLC GA Y NWLDC KASF EI N P TGP N Q P -IV EKCN V -- II I TDPDK VKE----ELE-KA KEIYKKR DE KTKS IRE EDV D VFY GCV M CQ SF APT H VC IIT PD RPSLC GSI NYLDA RAAA KI D P NGP I F E

ED G RV EV IGP EIK EIE - - GQ-SLP FGFWIRYY GKE LTE DYLDL L TRWTYFALEEGEG W ML LNTRD T IW LRL HK KNADK -HD-FE-HLGQAM I N L C K I Q F P K D G EV SL IGP DLA DME -E GG-KYAYGM I Y RV Y GEQ IEK DL EAV I ERR NHE Y Q SY IQG Y MH LN QRG G IW VRI SK DAVKK GLKSLK-Q IGRA T I M L FK AE L S ED M KV TL IGP DLD EME -E GQ-AYP YAM I Y YI A GEM VET DL EPV I ERR NHD F QNY IEG Y MH LN QRY D IW IRI GK NAIKK GLKSLI-Q IAKA T M M L YK NE L P ED D KV TI VGP DLK EME -E GK-TYP WAM I F NI G GEL VEA DL ESV V ERR VHD F VNY CQG I MH LN QRY D VW MRV SK DTAAK MDS-FD-SFGKAV M M L F K TE L P ED G KV TI VGP DLK DME -E GK-TYP WAM I F HV G GEL VEP DL ESV I ERR VHD F INY CQG I MH LN QRY D VW MRI SK DTAAK MDS-FE-PFGKAV M M L F K TE L P ED D KV TI VGP DLK EME -E GK-TYP WAM I F NI G GEL VEP DL ESV V ERR VHD F INY CQG I MH LN QRY D VW MRV SK DTAAK MDS-FE-PFGQAV M M L F K TE L P ED G KF TL IGP DIS EME -E GS-RHP FAM I Y KI A GEL VEE DL ESI V ERR NHD F QNY IQG L MH LN QRY D VW IRV SK DAVAK GLTSFE-P IAQAV M M L F K NE L P TD G KI SI LGP DLA DLE -E GK-SYP FGI L V EV T GEK LEE DL EGV F ERR IHE Y LNY IQG V MH LN QRY D LW MRV GK KAFAK GLN-SFHQ IGSAL I S L Y R SE L P EPD KV EV IGP DID EME -E GG-RYP FAIYVKAA GEE LEE DV EGV L ERR IHE F CNY VEG F MH LN QRD Q IW CRV SK NVTEK GFR-LE-H LGIAL R E L Y K EE F G ED G KI EV IGP ELK DME -E GK-KYP FGI L M EV S GEK LEK DL EGV I ERR IHE I CNY VQG F MH LN QRD K IW CRI GK DSHAK GFE-LK-H LGQAL S T L F K DE F P ED G KV EV KGP EID EME - Q GQ-VYP FAI N V EV A GSE LEE EL ESV I ERR LHE L CNY VKG F MH LN QRD Q IW CRVSTEAKDA GFR-LE-H LGKAL S V L F R EE F P E-D KV EI IGK DID EME -E GS-RNP FAI I V EV S GSN LEE DL EGV L ERR IHE F LNY IEG V MH LN QRD Q VW IRI NK DSFNK GLR-LK-H IGKVV Q R L F K AE F P ED H KI NV IGP NID DFE GEPK-RIP LAI I V EV A GKN MQL DF EPV L ERR IHY F MNY TEG V MH VG QRD T TW IRI SK DTYAS GFR-LE-H IGEVL Y A K M M DE F S ED H KI EI IGP NID EVD AD GVLRLP LAV I V KI A GKN MQE DF EPV L ERR FHY F LNY IEG V MH VG QRD M AW VRI SK DAFDK GFR-LE-H IGEVL Y A K M L DE F E ED G KV EV IGP DIDSVE - P GG-RLP IGI V V DI Y GRK MQE DF EPV L ERR IHY F TNY GEG F WH TA QRD L TW VRI SK EAFAK GAR-LK-H LGQLL Y A K F K QE F P TD G KI EV IGP DIDQI P -E GS-KLP LGI L V DI Y GRK MQA DF EGV L ERR IHD F INY GEG L WH TG QRN I NW LRV SK DAVAK GF R - F K - N YGEIL V A K M K EE F P VD G QV TV IGP EIDTV P -E GS-RLP LGI K V DI Y GRK MQE DF EGV L ERR IHY F TNY GEG V WH VA QRD L CW VRI SK DARAK GFL-MK-H IGELL L A K F K QE F P TD G KI EL FGQ DVS DM P -E GS-RLP LAIVVEAA GHK MNE DY EPI L ERQ IHH L LNY AQG V MH IG QRD I AW LRI GK SAAAK GFS-LK-H IGSIL H A K L H QD F G ED G RV EV IGP DIK DVK-P GS-RLP LAV T V QI G GRK MQE DY EPI L ERQ IHH L VNY AQG L MH IG QRD I AW YRI SK QAVDK GFT-LE-H IGKIL H A K F H QD F G E-DKV EI IGK DID EME -E GS-RNP FAI I V EV S GSN LEE DL EGV L ERR IHE F LNY IEG V MH LN QRD Q VW IRI NK NSFNK GLR-LK-H IGEVV K Q L F K E H F P

----PKGKVYHVPTN-------PDD------------------------FFTGISYDISP RQAGQ R VR KHD MY CELGG PK - Q R Y S S F F FI E V V K G - E D KI -------------------------------------------------EVEKF PF D I NP MY EGE R VR KAG LY AELGG QS-K---PGFELV L F E P E P E NI -------------------------------------------------EEAEF PF D I SP MY EGE R IR KGD MY VELGG PT-Q---PGFELV M A L P - M D QV --------------------------------------------------MAEF PF E I SP MF EGE R VR KDG MF VELGG PK- S---MGL ELV R A K D - L D EI --------------------------------------------------MSEF PF E I SP MF EGE R VR KEG MF VELGG PK- S---LGL ELV R A K P - M D EI --------------------------------------------------MVEF PF E I SP MF EGE R VR KEG MF VELGG PK- S---LGL ELV R A K P - M D EI --------------------------------------------------ADEY PF E I SP MF EGE R IR KDG MH VELSG PK- S---KGY ELV R A T S - M D EI --------------------------------------------------FEDI PVEVGLVH EGE R IR KED MR VELGG PKV S---EKF ELV L V K K - S D EI -------------------------------------------------SLADL PV D V SP RH EGE R IR SGD MY VELAG PK- S---FGA EL F K V V D - P D EI -------------------------------------------------MFGDI PV E I SP MY EGE R IR AAN MF VELAG PK- S---VGA ELV L V S - - - D DV --------------------------------------------------FEDI PV D V SP MH EGE R IR SAN MF VELAG PK- S---IGA ELV Q V K - - - D EV -------------------------------------------------MFDDI PV S V GP MN EGE R VR GPD MY VELAG PK- S---YGF ELV K V V N - - - K A ALGFPVITDQIVTE-------VPTLLLTQKDFDKIPATSLEARNIKIKVTEIDIPVGFAAAF EGE R VR KDN LY AEFGG SK- T---ECW ELV R S K E - L S EI ALGFPVITDQTVLE-------VPMNLLTQKDYDKIVATSLEARGIKIKVTEIPIPVSFAAAF EGE R IR KSD MF AEFGG NR- T---EAW ELV V K K E - A T EV FTGFPVITDQPLPEDKQ----IKDWFISEPDYDKIVQTALEVRGIKITSIDIDLPI N F GP AF EGE S IR KGD MH VEFGG GK- T---PSF ELV R M V G - P D EI FTGFPVITDQPLPEDKQ----IPDWFFSVEDYDKIVQIAMETRGIKLTKIKLDLPI N F GP AF EGE S IR KGD MY VEMGG NR- T---PAF ELV R T V S - E S EI FMGFPVLTDQELGEDMQ----IPDWYLSEPDYEKIVPLALEVRGIKLTNIEIPIPV N F GP AF EGE T IR KGDTY VEFGG GR- T---TAF ELV S M V G - P D EV NFGFPIIADKSVLPILPG-VESVDSVLAGIPYSRIVSKAMDARGIKVNVTKVPVPV A Y GP AY EGE R VR GEQ IY LECGG GR- T---QMV ELV T T A K - M D DV NWGFPTIADTDIPEILPTGVCTYEHVVSSVPYDKMTARSIEVRGLKLTVAEIPVPV A Y GP AF EGE R VR RDD VY LECGG GK- T---PCC ELV Q I A D - M N TI KAGVPVITNNEVPVI--------KGALESSDIDNIVENALKMKGVKVKVVEFDIPV S V GP MN EGE R VR GPD MY VELAG PK- S---YGF ELV K V V N - - - K A

ACETYL-CoA SYNTHASES AND Ni-CONTAINING CODH 403

Appendix 3. (Continued)

Met. Ions Life Sci. 2, 357–416 (2007)

Met. Ions Life Sci. 2, 357–416 (2007)

379 342 35 6 333 333 333 33 5 33 6 33 7 34 8 33 2 331 62 9 62 9 65 3 650 64 7 655 65 8 613

454 428 44 7 411 411 411 41 3 41 3 41 5 42 6 41 0 409

A NME3 ANME1 A.fulgidus Ms.bark Ms.thermo Ms.acet M.burtonii M.hutchins M.kandleri Mc.maripal M.thermaut Mc.jann1 A .metallir C.difficil C.hydrogen Mo.thermo D.hafniens D .ethenog S.fumaroxi Mc.jann2

ANME3 ANME1 A.fulgidus Ms.bark Ms.thermo Ms.acet M.burtonii M.hutchins M.kandleri Mc.maripal M.thermaut Mc.jann1 A.metallir C.difficil C.hydrogen Mo.thermo D.hafniens D.ethenog S.fumaroxi Mc.jann2

RTPR SRK FVR ADG GW N RI VWI PK E YK QIL T E FIPA--------- EMF DKI A TEEDCIDP EELKEF L KRVHH PV VTLWKNGET------- - - - - - - P D - - QYF Y SSK FI Q LDG GW R RV VWM A S DLK ERV V D AISD--------- EMK DKI A TEKDVKDI EELREF L QRVDH PV VKGVIREVDGERVTEG W A V K E E - - - - AYF R SKK FIQ ADG GW Y RT VWM PK E LK ERV A K YIPD--------- DIR DKI A TEEDAKTL DELREF L KKVDH PV VKGVVRPVDGKKITNG W V E E E E E E A E E N YF Y SPK FIQ ADG GW N RV VWL PA M LK EKI A E TIPE--------- DIK DKI A TENDATDI ESLKAF L QEKNH PV VANWASEEE------- - - - - - - E E E E E NYF Y SPK FIQ ADG GW N RV VWL PS M LK EKI D E AIPD--------- DMK DKI A TEKDVTDI ESLKTF L KEKNH PV VANWAAEAE------- - - - - - - E E E E E N YF Y SPK FIQ ADG GW N RV VWL PS M LK DKI I D TIPE--------- DLK DKI A TENDSTDI ESLKAF L QEKGH PV VATWAAEEE------- - - - - - - E E E E E NYF R SPK FIQ ADG GW D RV VWM PK H LK DRV L A DIPA---------NIADKV A TEEDASDL DSLKNF L TEKDH PI VQRWEEEAE------- - - - - - - E E P E A EYM R SVK FLQ ADG GW N AI VWM PA E IK EML S G IIPE--------- DVAGA I A TEQDALDINALKSF L TSHHH PV VSKWIVPED------- - - - - - - T E E E GYM E SDK FL Q YDG GW E RV VWM PK A LK ERM K H AIPD--------- ELY DKI A TEEDATTV EELREF L EKVEH PV VERWAEEEE------- - - - - - - E E E E K EYM R SPK FLQ ADG GY E RI VWL PK E IK EKV L E SIPE--------- ELR DKI A TEEDVSSIHNLKKF L NEKDH PV LKRIAELDA------- - - - - - - P F E E Q EYM R SPK FLQ ADG GY H RV IWM PR E LK ESV L E FIPE--------- DVR DKI A TEEDATSIKDLRRF L RDNEH PV LERAAVEET------- - - - - - - E P E E E SYM K SPK FLQ GDG GW E RV VWL PK E LK ERV K D AIPE--------- ELY DKI A TEEDVKTT DELIKF L KEKGH PCAERIGAEVE-------- - - - - - E E A I E YFI S SKK FIS AEG GP G RI VWM PK E LK DHV K D RLDQTAKEMYGIDNFTDMI C DETIATDS EEVLNF L TQKGH PA LTM--EALM------- - - - - - - - - - - HFI S SKK FAY AEG GP E RI VWM PK E LK DYV A D KLNATVKEMTGIENFCDMV C DETIADDS EGVLAF L EEKGH PA LAM--ESVM------- - - - - - - - - - - SYI G SRK FVK ADG GL A RV VWM PK D LK EQL R S IIEERAEEEGLGR DFI DKI A DETVGTTV DEVLPF L EEKGH PA LSM--EPLL------- - - - - - - - - - - TYI V SKK FIS ADG GI A RI VWM PK S LK DFLHDEFVRRSVEEGLGE DFI DKI A DETIGTTV DEILPY L EEKGH PA LTM--DPIM------- - - - - - - - - - - SYI I SRK FIP ADG GI S RI VWM PK E LK EFL K E DLVARSIDEGLGE DFI DKI A DESVGTTV EEIIPF L EENGH PCFSL--DPLM-------- - - - - - - - - - YNITQRK FIS AE DS LL RL VWM PK S LK EEIRERFNARAAELGV-P DLL ERI A DESVGTTE EEILPF L EEKQH PA LTL--EPLI------- - - - - - - - - - - FNVTQRK FVS GDG GI H RL VWM PK I LK EEI R D RLEKRGQDIGI-PNFVDMI A TEENGVTE EEILAF L EKMGH PA VTM--DPIL------- - - - - - - - - - - SYM K SPK FLQ GDG GW E RV VWL PK E LK ERV K D AIPE--------- ELY DKI A TEEDVKTT DELIKF L KEKGH PI VKKTEEEVVEEVEEEK - - - - - - - - - - E

M PK G ECV DP E GGE YTG IN EKIYEKSNRTYKRV VM YSSITY PQ TN CGC FEAAIFY IPD V D GLGLVD R R-YGG ET PLGLT FSK LAGL I SGGQQ NH GY CGI SV I PK G DCI DP I GGE YTG VN EAAVRLSSGEYDSI KI HSFLEK PH TS CGC FEVAGFY IPEV E GIGWVH R GSKVSET PIGLP FST IAGS V GGGKQ VQ GFLGI G I V PK G ELL DP I GGE YSG VN EFAKQESGGEYERI KL HSFFEY PH TS CGC FEV IGFYMPEV D GIGWVH R G-YAEPA PNGLT FST MAGQ T GGGKQ VV GFLGI G I I EK G ELI N A D TGE YSG VN EIAKKLSSGEYDKI NL HSFFEY PH TS CGC FEV VGFYIPEV D GIGWVN R E-YQG MA PNGLG FST MAGQ T GGGKQ VV GFLGI GV I EK G ELL DA K TGE YSG VN EVAKKLSSGEFDKI KL HSFFDA PH TS CGC FED VGFYIPEV D GIGWVN R E-YQG MA PNGLG FST MAGQ T GGGKQ IV GFLGI G I I AK G DLL DA N TGE YTG VN DIAKKLSAGEFDKI KL HSFFDS PH TS CGC FEV VGFYIPEV D GIGWVN R E-YQG MA PNGIG FST MAGQ T GGGKQ IV GFLGI G I I PK G DTI N A V AGE YSG VN DLAKSLSSGEYDRI NL HSFFEY PH TS CGC FEV VGFYIPEV D GIGWVD R D-FTG VA PNGLP FST MAGQ T GGGKQ VV GFLGI G I I AK G EYL DS E HGE YSG VN EHAIKRSMGAVNRV WL YSAFGY PH TS CGC FEA IAFYIPEI E GFGIVH R R-FAG PTVNGLA FST LAD S T AGGRQ VE GF HGI S L I EK G ECL DP E AGE YEG VN EAVKEHSQGTVERV YL HSCLEY PH TS CGC FQA VVFYIPEV D GFGIVD R E-YPG ET PIGLP FST MAGE A SGGEQ QP GFVGV S Y I PK G DLL DP V SGE YTT LN ATVADKSQSTFDKV YL HSIFGH PH TS CGC FEA VAFYIPEV D GIGIVH R D-FKG DT PMGIP FSA MAGQ C SGGKQ VE GF AGL CV I EK G EVL DP E RGE YAN VN AAVEENSQGTTDRV YL HSVFGY PH TS CGC FEA VAFYIPEL D GIGIVN R D-FRG ET PLGIP FSA MAGQ C SGGKQ VE GF SGL S L I PK G ECL DE K LGI YSG VN EVVRERSQGTVEEV TL HSALEK PC TS CGC FEA IVFYIPEV D GFGIAH R G-YKG ET PMGIP FST LAGQ C SGGKQ VP GFVGI S I I AK EG VEDE V KGI WSA VN EHVDEKSQGAVSRV TL YSLLED PM TS CGC FEC IAGIMPEA N GVVIVN R E-FPSTT PVGMT FGE LAS M T GGGVQ TP GFMG H G R I EK G ECL DD R TGV WNS VN ETVNQISQGAVESV TL YSILED PM TS CGC FEC ICGIMPEA N GFVVVN R E-FASVT PVGMT FGE LAS M T GGGVQ TP GFMG H G R I PK EG LI DP V KGQ W E S FN EYIYKNSQRTIERM NL YTIMEY PM TS CGC FEA IMAYLPEL N GFMIVN R E-HSG MT PIGMT FST LAGM V GGGTQ TP GFMGI G K I PK EG EI DP I KGI WKS VN DYLYTASNRNLEQV CL YTLMEN PM TS CGC FEA IMAILPEC N GIMITT R D-HAG MT PSGMT FST LAGM I GGGTQ TP GFMGI G R I PK G LAI DE V KGM WQS VN DYLRPSSNNTLEEV NL YTLMDR PM TS CGC FEA IMAIVPEA N GLMITT R E-HSG MT PCGMT FST LAGT V GGGLQ TP GFMGI G R V NK G AVI DA K LGQ WSG IN QFVEKTSGGVVSHYNL YSIIYD PM TT CGC CEC ISAILPL C N GVMTVN R E-HTG MT PCGMK FTT LAGT I GGGI T T P GFLG H S K V QK G ETL DE T LGN WKG VN DFVYKASRQSISSYNFYSLVFD PM TT CGC CEC IAAVLPL C N GIMTVN R D-YTG MT PTGMK FTT LAGV I GGGN V T P GFVG H S K I PK G ECL DE K LGI YTG VN EVVRERSQGSVEEM AL HSALTN PC TS CGC FEA IVFYIPEV D GFGVAH R N-FRG ET PFGLP FST LAGQ C SGGKQ VP GFVGI S I

---------------------------PLNLPPPN----------------TDWDQEEEKKARGKGKNMLVIPP----P-------AA--EVEAEAVPSAA---------------QVPLASIPMM-ASGAVAGG-GTPIRIILKDADISIGKIIVKKKG-------GK-VAE--EAAAEAAPAAQPAQAAQPMAMQPMPMQMPGFQLPAL-QMPAASAA-PAGVKLVIKDAKITIEKVIIKKAEKEKKG--GK-EEEEEEAAVSAA---------------PMMMPSAGFQMPAMASMPMMSGG-TGGIKLTFKNAKISIDKMIVSEKK-ENK------EGEEEEVAAEAA---------------PMMMPAAGFQMPAMPAMPMMSGG-AGGIKLTFKNAKITIDRMIISEKK-EKK------EEEEE-VAVAAA---------------PMMMPAAGFQMPAMP---MMSGGSSGGIKLTFKNAKITIDKMIISEKK-EKK------EEEVQEATMFAA---------------PPTMQNT-MPMQGMPMMMPSSSG-TGGVKIILKNAKVSIEKVIIQKKE-----------ISSD-----FGPAT------------------PVMSFGELP-L--M----AGGVKIILKNAKIYAERVIIQPADTHRKKQGGKND APEEE------APAEE-----------------PTMEVKELP-I--APGG-GLNVKIVLKNAKIYAEKVIIKRADREDKS-----VVETE-----AEPAGE---------------FVQFAQIPEMA-I--PTS---GGIKIILKNAKIYAEKIIIKKK-----------EVEEAYP--EETPIPE---------------GVPVMAAPEMT-L--PAA---GGFRIVLKNAKIYAEKVIIKRK-----------EEEVEEEMEEVEGIE-----------------VPTMTLPGTF-A--G--L-PPGIKIVLYNAVIKAEKIIITKEEPEKKKKKKK-- -- -- -- -- -- -- -- -- -- ------------------------------------------------------------------- -- -- -- -- -- -- -- -- -- ------------------------------------------------------------------- -- -- -- -- -- -- -- -- -- ------------------------------------------------------------------- -- -- -- -- -- -- -- -- -- ------------------------------------------------------------------- -- -- -- -- -- -- -- -- -- ------------------------------------------------------------------- -- -- -- -- -- -- -- -- -- ------------------------------------------------------------------73 7 ------------------------------------G------------------------------------------------694 -EV--KATEEEKE---------------------GIEVGEL---ITKLA-KEGGIQIIMKNVKIVINLNVKR--------------

280 242 25 7 234 234 234 23 6 23 7 23 8 24 9 2 33 232 53 0 53 0 55 4 551 54 8 556 55 9 514




1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

83 80 83 83 83 83 83 88 88 88 93 89 89 89 96 89 84 83 83

182 179 181 182 182 182 182 187 187 187 191 187 187 188 195 187 183 181 181

M o . t h e r mo D.hafniens C.hydrogen A.metallir C.difficil D.ethenog S.fumaroxi ANME1 ANME2 ANME3 A.fulgidus Ms.thermo Ms.acetiv M.burtonii M.hutchins M.jann M.kandleri Mc.maripal M.thermaut

Mo.thermo D.hafniens C.hydrogen A.metallir C.difficil D.ethenog S.fumaroxi ANME1 ANME2 ANME3 A.fulgidus Ms.thermo Ms.acetiv M.burtonii M.hutchins M.jann M.kandleri Mc.maripal M.thermaut


L Y A A TGA N Y E AM T ALA K E N N C P L A - V Y G N - GL E E L AE L V D K I V A L GH K Q L V L DP G A - ---RETSRAIADFTQI R RLAIKKRFRSFG YPI IALTTA-ANPI G S A TEA N Y E AV V NLA K E H N V P V I I - K A D - GL D A L AA L V E N A Q K L GY K E F V L DP G A - ---RTPSQTLANLTHCR RLAIKKKFRPFG YPV IAFTSK-TEPL Y A A TAD N Y E QM V ELA K K Y N V P L T - V S A K - GL D A L AE L V Q K I T A L GY K N L I L DP Q P - ---ENISEGLFYQTQI R RLAIKKLFRPFG YPT IAFALD-ENPV Y G A TKD N Y E AM V SLV K D Q K L P L G L - K A D - GL E E L YN L V E K V Q G L D Y KE L V L G T E A - ---SSVKE TFKNAVQI R RTALKEQDRKFG YPS VVFANELANNV Y G A TKE N Y K DM I EV V K G A S LP L G - V K A G - SL E E L Y E T V E L I Q A AGY K E L V L DV T G - ---ENIKD TYTNAIQV R RTALKEQDRTFG YPS IVFANRLSN-L Y A A TAE N S S EM A ELA K K Y N C P L V - A K A A - NL A D L AD L S V K L Q S L GI K D I V L DS S A - ---ENLKQALSDNVAI R RSAILKKFRPMG FPV INFPGQMTQ-I Y A A DAS N V D AL G ALA K E V A C P L A - V K G D G SL E S VIA L T E K L S G M GI K D L V I DT G S - ---RELKKVFEEQILI R RSALKNRFRPLG YPT MIVANEMA--I Y A A TTE N W R DV A ELA L T H N V P V A L - S A P N D PD A L KS L A L T F A S M GI T D L V L DP G T F PT-DGLRD SFENFLKI R RAGING-QTDIA YPL LAVPMTVWERL Y A A DKT N W Q AI L ELA K Q Y N V P V T L - F S - P DL D T L DR L A T T F A A A GI D D L V L DP G T Y PTGKGLAD TFSRFVRI R RAGIVDGNKGIA YPL MAVPMTAWMTN I Y A A TED N W K P F L ELA T R Y E V P V T L - F S - A DL D M L NT L A V T F S S A GV T N I V L DP G T Y PTGKGLEE TFSRFIRI R RAGIVDGNKGIA YPL MSVPMTAWMVN M Y A A TEG N W K E F L KLA L E Y K V P V T L - R A - K DL D L L KS M A V T F K Q A GV K D I V L DP V T E PLGEGLKG TFERVIQL R RTAIAGQDKDVA YPI MITPIAAWLII Y A A NKD N W K EV G ELA L E Y N V P V V - V S A F N DL D G L KT L A K T F A E A GI K D I V L DP G T Y PSGQGLKD SFTNFLKI R RAGIMG-DTEIA YPI MALPITAWMAL Y A A NKD N W K EV G ELA I E Y K V P V V - V S A F N DL D A L KT L A K T Y A E A GI K D I V L DP G T Y PTGKGLKD TFTNFLKI R RAGIMG-DTEIA YPI MALPFTAWMAL Y A A NKD N W Q EV A ALA Q E F N V P V T - V F A P N DL D L L KS M A K T F A D N AIE D V V L DP G T F PTGKGLKT TLNNFLKI R RASING-DRDIA FPI MAVPFTAWMAI F A ITKE N W K EM G ELV L D T G C P V V - V S A P G DL S L L RT L V R T L Q E W GI V D M V L DP G T F HS-DQIGQ TIHLFTAL R KAAVKNFDSLSG YPI LGTPISVWNGV Y A A T K E T L N D F I KV I K E V K K DV V L V L S S N NV K D L KN M A A K C L A N GI E D L V L EP H T Y --PENIAE TLDLNVMI R RSAIEKEDKYLG FPI LNLPINAYYYA L Y P A TEE N V E DL A KLA A D G D C P L G L - H A - R DV E D L VP L V V E A Q Q Y - T DD L L L DP G T E FGPHDVVS TTDKLAEI R KAAIEE-FESFG YPT LVTTFPYAFLI Y A A TEN N I D EM S ELA L K H N C P I S L - F V P N DL E K MKQ L S R K L R E S GV K D I V L DP G T Y -IGEAIGD TMDNFIMI R RLAVEEKDDDFR FPI LAVPAVCWINL Y A A RED N L G EM A ELS V S Y G C P L V L - F S P G DL E E MKN L S R R L R S L GV T E I V L DP G T F -TGEGIGD TIDNFVMI R RLAVEDGDDDFR FPI MGIPALSRLT-

R H D K R FY H E T A I A I QV SD N L S S E E LKAKVEA I N G L N F D RV GQ H Y TIQA I A I R H D A DD P A AFKAA V A SVA-AATQLNLV LMADDPDV LKEALAGVA DR KPL R H D K T FY H P T T L L I EI AD T L SD E E VQGKLQE I E G L E F D RV GL H Y TID G V A VIEA S G S PEQ FAKV V A QVA-AGTERSLL LLSDNADA LKAALPGVAGR KPL R H D K R FE H P C G L A I LV ED T L SE G E IKERVEK I N K L V F D RV GQ M H SVNL V A L KGS S QD A A TFAKA VATARE-VTDLPFI LIG-TPEQ LAAALETEGANNPL R H E K T FV N R N R F A V M F T D Q L SE E A IQSKLDN I K K V N Y V RI GE T M K VE I A A I KYV S -D K E KYFNL I K KVKESGISIAYM LICEDVDV LKEALELVK DE KPI R H E K T L V S R N R Y A V S F C T CM SD E A VDAKIAN M K K V D Y V RI GE Q M K VE M A V L E Y C G -D K D AYLKL I D KIKGSGLEVAYI LACDDAQV VKEAVEVLK DA RPM R H E K R FE H Q P G I A L LI SD S L SE T E Q D AK L A A F T R F Q Y K R V GA I L K PD M L A L RAD S G S GEK YLKL VKLAAG-KCPASLM LMCADTAV LDESLKICA ER KPL R H E K T FV N P P G F A A VV G T D E D D A S VSGK I E R F R Q C Q F D R V GL K L R PE L F A V R D A G GD A G KFQGLAKKISD-ETGGSLI LMSGNVDA LKAAAGALK EK KPL R H K L T FF N K T A F A Y DV WD T M SE E E LVGRVNA I Q N Y K K F YI GE F L T LD A V A I RCA S GD A D QFAQTTAKVVE-TTELPLL LCSLDPEV LGAGLAVAA DR KPL R H E L T FY D R T S L A Y DV WD T M AE P E LRERVKG I T K W R K F YV GE F L T LD A I A I RSV S CE A E TFARC V K CVAE-ETDMPLI LCSFDANV LEAGLKEVS DRNPL R H E L T FF N R T A L A Y DV WD T M KE E E LRERVRE I T N W R K F YI GE F L G VD A I A V RSV S GE A K TFAQC V K RVAE-ETDLPLV LCSFDAKV IEEGVKEVS DRNPL R H E L T FF N P T G F F Y DV WD T M DD K A LEERCDR V V S Y K K F YV GN F L T LD G F A V RCT S GD P K RYREV V K KVA-SY-GKPLI LVALDSEC MKAALEEVA DQ RPL R H K L T FF N K T K M F Y DV TD T M DE A A LLERTKK V A D F R K F YV GR N L L LD G V A I RSV S ND P E KFAAA V K KVSE-V-GIPMI LCSFNPAV LKAGLEVAK DKNPL R H K L T FF N K T K M F F DV AD N M DE A A LVERVNS I A N F R K F YV GR N L L LD G V A I RAV S ND P A KFAAA V K KVAE-A-GLPMIFCSFNPAV LKAGLEAAK DL KPL R H K L T FF N K T A L A Y DV WD T M EE T D LVERVNK I Q D F K K F YV GD Y L T LD M I A V RSV S ND P E KFAAT V K KVME-TTEFPMI LCSFEPAV LRAGLEVAA EK RPL R H E F T Y Q N P P P I A I DI SD D M E N E Q IAIRIKK V Q D F S Y T YI GR P L S LD A L A V RSV T Q N PYR YTEV I Q HICE-LCNLPLI LCSTDPNI IKAGLSACA ER KPL R Y Q L S FF N P T P I G V DI SD E L SE E E IKNRAKE I E N F V F E R TGE K L K LD F I V I RNA S GD V E KFKKA I E IV-EKETKMPIC IASLNPEV IKEALKVVK-S KPM R H E L S FF N P P P V F V TV YD D M EE D E IAGKTEE I Q E F Q V E RV GE V L K LD G V A VVSR T GD P E KYARA VEIAVERSEDLAVA LITTDPKV MEAGLDVF- DE RPL R Y E L T Y Y N P T A V A V DL SD D M DD S A F D QKLVK I K N L E F E R TGE I L K LNA V A L RNK S GD A E KFKKAAEKL-KDSE-IPVI LCSFDPKA MDAALDVIGSK RPL R Y E L T Y Y N P T A L V V DL PD D L P S E E LLNRAQR I M E L E F E R TGE K L T LD A I A L RNR S G S PEK FAEAAEAISK-L-NFPVV LCTFDVEA MKAALEVLG DQ KPL

M - - - - - - - - - P L T GLE IY K Q LP K K N CGE C GT P T C L AF A M N L A S G K A S L D SC P YV S D A A---------REAL DAAAAPP IAK VVLGA GPTA VEM GD ETELF M - - - - - - - - - A L T GLE IY K Q LP K K N CGE C GT P T C L AF A M A L A S G K G S L D AC P YV T D E A---------REAL DSASAPP IKA IKFGN GS-- V-L GD ET VLF M - - - - - - - - - G L T GLE IY K H LP K K N CKE C GQ P T C L AF A M Q I A A G K A G L D AC P YV S D E A---------KELL ESASAPP VAL IKVGK GEKV LEI GH ET VLF M - - - - - - - - - A L T GLD IF K L TP K K N CKD C GF P T C L AF S M K V A S G G S E I E KC P HM S P E A---------LEKL SEATAPP MKA LKFGK GDNE YTI GG ET VLF M - - - - - - - - - A L K ALD IF K L TP K K N CKD C GF P T C M AF S M K V A S G A V E V G KC P HM S D D A---------IAKL SEATAPL MKA LKVGA GASE YEL GG ET VLF M - - - - - - - - - S P T GIE IY K F LP R T N CGE C GV P T C L AF A M S L A G G R A E L S AC P YV S E E A---------KQKL EEASAPP VRT VSIGT GDYSFKT GG ET VMH M - - - - - - - - - A L T GI Q IF K L LP K T N CGE C GV P T C L AF A M N L A S G K A E L E KC P YV S E E A---------KETL ASESAPP IRP LTIGT GDYA VKL GG ET VMF M - - - - - - - - K I N S PL Q AY E Y LP K T N CGD C GE P T C M AF A S H L I D R S M K V E EC P PL L A E E-----YNDNYSTL SDLLAPE IRE VVIGT GDHA ITI GG DD VLY M - - - - - - - - K L S S PMD VW K Y LP G T N CKE C GE K T C L AF A S L L L E R N K K L E EC T PL L E - ----PKYASKAKEL AELLAPE IRE VAIGV GARV SKI GG ED IMH M - - - - - - - - K I S S PME AW K Y LP G T N CKE C GE K T C L AF A S L L L E R K K K L E EC P LL F E - ----PKYAEKGKKL AELLAPE VRE VEIGI GDKA IKI GG ED IMH M - - - - - - - - K V K S PLE VY N Y LP R T N CGE C GF D T C M SF A A H I L D R S V T P L DC K PL V R D AEKDPKVKKKLEEL LELTAPE IAE VVIGV GENA VKI GG EE VLH M - - - - - - - - K I N S PLE AY K Y LP Q T N CGE C GE A T C M AF A S K L I D R S G K P T QC P PL V K E K----KFAKKLAEL ERLLAPE IRE ITIGV GDRA VKI GG DD VLY M - - - - - - - - K I N S PLE AY K Y LP Q T N CGE C GE A T C M AF A S K L I D R S G K T S DC P PL I K E K----KFAKKLAEL DRLLAPE IRQ VTIGV GEKA VNI GG DD VLY M - - - - - - - - K I N S PLE AY K F LP G T N CGE C GE T S C M AF A S H L I D R S L K A T DC T PL V N E S----KYKKKYDEL VTLLAPE IRE VVIGV GDNA ISI GG DD VLH M K E E H K K S I R E I S PID VY K L LP R T N CAE C GE A N C M AF A T K V V N G E A F I E GC P PV L T K N-----YEKDLLKL QELLAPP VRV IHFGT GNNQ LKI GGKH VLY M - - - - - - - P K K I S AMD IY K L LP K T N CK KC GY P S C M AF A T K L L E K E A T I D QC P IL N T P K-----FEKNKKKIIELIS PP VKE VWFGNEEKK AVM GG DE VMY M - - - - - - - - A Q L S AMD VY N L LP K A N CG AC GC K T C M EF A T K L V N R E A K P E DC P KL D D E S---------LEKL QELLAPP VKE LTIGE GDRE VTV GG DE VMF M - - - - - - - - - K V T AMD IY K L LP K T N CG KC GE A S C M AF A A K L S Q K E A E L S AC P QL K G A D---------FEKL ANMLAPA VRE IKIGT GDRA VTI GG DE VLY M - - - - - - - - - Q V T AMD VY R L LP K T N CG KC N E A S C M AF A T K L I E K E V T L D DC P QL S G D E---------RQKL ENLLAPA VKE ITFGPEENQ VVV GG DE VLY


Appendix 4. Alignment of γ Subunit Protein Sequences

Met. Ions Life Sci. 2, 357–416 (2007)

Met. Ions Life Sci. 2, 357–416 (2007)

481 463

480

455 455

354 351 353 357 357 356 357 370 374 383 376 373 373 373 381 384 366 367 366



274 271 273 275 274 274 274 283 285 285 288 284 284 285 292 285 278 278 278


---------------------A------N----------------------------------------------------------------------------WKEKNQTA ----------------------

I E S T K IPS Y L LS V DTD GL S V L T A Y A DG K F EA E K IAAV M K K V D - - L D N KV K R H RI I IP G AV AVLK GK LE- D L TG W EVIV GPR EASGI V AF ARANLAS---V E A S K IPS Y I LP I DTD GT S V L T A Y A AG K F E P E K IADI L A K S G - - V G D KV N H R NL I IP G YV AVIS GK LQ-EISG W KVIV GPR ESSGI V SF TRAM------V E G A R IPA Y I LP V DTD GT S V L T A W A AG K F T P E K IAQF L K E S G - - I A E KV N H RKA I LP G GV AVLS GK LQ-ELSG W EILV GPR ESSGI N SFIKQRWNV---I E R S K VPV W V A I P DAG GY S V L T A W A AG K F GA G S ISKF I K E S G - - I A E K T K SR KL I LP G LV AVLK GE LEDELPEW EIII GTE EAMHI P KFLKQLNNTVEAN I E R S K VPV W L VIP DAG GY S V L T S W A AG K F GG N S ISAF I K E S K - - V E E V T N CK DL I IP G KV AVLK GD IED N L PG W NVVI GPE ESMEL P KFLKGYQEKACQT M E N S R VPC Y L MIK DTE GL S V M T A W A AG K F GA D T IGKF V K N S G - - I A A KI N H R KL I IP G YASMES GG LEE E L SG W EIMV GPR EAAHI P AYLKQFKP----I E S S R VPS W L CVQ DTE GL S V M T A W A AG K F SG E S VGTF I N K C G - - I K D KV K H K SI I IP G YA AGIS GE LEE E L GG W NVVV GPR EASQI S KFLRAFKPQ---L A S N D IDC Y L CV V DTD GI G V E A A V A GG Q L TA T K I K G M F D S A G F DV K E K T DH G SM I LP G LA ARLQ GD VED E - TGL T VMI GPP DSGQI P RWMESKWET---L S S S K IDC Y L MV V DAD GL G V E A A V A GG Q L TA D M IKDT V A S Y D - - A E K KV T H K TM V LP G LA ARIS GE TED A - TG W SVLV GPR DSGRI P GWMTDNWPPKT-L A S N K IDS Y L MV V DGE GM G V E S A V A GG Q L NA E T MKDA L E S F D - - V E R KV K H K IM V IP G LA ARLS GE TED V - TG W TVLV GPR DSGRI P GWMENNWPPETFS L T S G G IKG W L LV L NTE GL G V E V S V A GG Q F TA S K VKEL I E E T K - - I E E KV N H R YL V IP G LA ARLQ GA IED E - TG W KVLV GPM DSGRI K GWLEKNWPPKE-L S S N G ITC W L LA V DTD GI G V E A A A A GG Q L TA D K V K D A F E K S G F DL K K DV T H N TV I IP G LA ARLQ GD LED K L -GA R VLV GPM DSGRL P GW FEKNWPPK--I S S N G IDC W L LA V DTD GI G V E A A V A GG Q L TA D K V K D A F D K A G F DL K T AV N H N TV V TP G LA ARLQ GD LED K L -GA N VKV GPM DSGRI P GWMEKNWPPK--L S S N G VDC Y L TA I DTD GI G V E A A V A GG Q L TA A K IKKG L D D A G F D M E K L - TH K VI V LP G LA ARLQ GD VED E - TGA G VMI GPA DSGRL P GWMEQNWPPQK-I K A A K IDC Y L IV I DTG GL S V E A A V A GRYLTA A K IAES L K E W K - - A D T LV S H K NL I IP G LA ARLS GE TEE E - TG W RVLV GPR DSSGIGQM IRECWPPADEF E K D NVTC W L LV M DTG GK A V D V S V A GG Q Y NG E N A K K L I E E T G - - I A D KV S H R II I LP A L A ASTR GD IED K - TG W TCVV GTR DSSQV G DFLRNNWDKILKE L E S A D IDC W L LV I DTG GL A V D V S V A GG Q F TG E A VKEV I E E T D - - I E D KV E H R VL V IP G KA AAVK GD VED A - TG W DVMI GTQ DSSEL P EFLEKEGLLRIEE L K S G K VNC Y L LV L DTN GK A V D V A V A GG Q F NG K A A A E L M K E T G - - I E K LV N H K KL I IP G LA ASVS GD IED Q - SG W EVIV GTR DSSEV P AFLAKIW-----L K S G D VTA Y L LV L DTE GR A V D V S L A GG Q L NG P A VADL I K E T G - - I E E RV R D K VM I IP G LA APAS GE IED D - TG W RVLV GPR DSSGI P DYLDKLASE----

- - - - - - - - - - - - - - - - - - - L DE V L Q AV N Y V T K Y A S LV V L R T D A K E HL L P L L S WR Q NL Y T D PQV P IR VEEKLNE IGAVNE-N SPVYV TTNFS LT YYSVEG E - - - - - - - - - - - - - - - - - - - L AE I T E AS V Y V A K Y A S AI V L K A S A K A HI L P L M A LR Q NL Y T D PQK P IQ VEPILHT VGEVNE-N SPIYI TTNFS LT YYSVEG E - - - - - - - - - - - - - - - - - - - Y Q A V M E AS V Y I A K Y A G II V L N T V E P A DI L P L I T LR L NI Y T D PQK P IA VEPKVYE ILNPGP-DA PVFI TTNFS LT YFCVAG D - - - - - - - - - - - - - - - - - D S N ME I A I AS L F V V K Y G S II V L E D I D Y A K AL S L F S LR Q NV F T D PQR P MR VEPDIYP INGADE-N SPVLV TVD FA LT YFIVAG E - - - - - - - - - - - - - - - - S N P M ME V A L S S I F T IK Y G S II V I D D I S Y A K AL P L F A LR Q NI Y T D PQR P MR VEPKIYP INNPDE-N SPVLV TVD FA LT YFIVAG D - - - - - - - - - - - - - - - - - D P M KE S L F AG I L T A K Y A G IL V L S D F S G E TL F P L L V TR M NL Y T D PQR P LATTEGIYE INNPDA-N SPVIL TCNFS LT YFIVSG E - - - - - - - - - - - - - - - - G D L M EE T V I AA A F V A K Y G A LM V M S D F Q P H SV F P L L L ER L NI Y T D PQR P M T AEQGIYP INNPDE-N SPVLV TCNFS LT YFIVSG E - - - - - - - - - - - - S D G I S A A Y WE S V L AS I F T V R Y G D LM I M H S I E P Y SL L P E L H IR D TV Y T D PRK P VM VDAGVFE VGSPDR-K SPVFV TANFA LT YYTVES D - - - - - - - - - - G N Q D A L D T S Y WE A I I AD T L I L R Y A D IM I L H S I E P H SL I T E R T L V A NI Y T D PRR P VS VDPGLRE VGSPTD-K SPLFI TTNFA LT YYTVES D - G N - G T G T G A K A E N T L E T S Y WE A I L AD L F I L K Y A D IM I L H S T E P H SL I P E R T L V A NI Y T D PRR P VS VEPGLRE VGTPTAGE SPLFV TTNFA LT YYTVES D - - - - - - - - - - - E G D D V T K S Y WE A V I AS I F I V K Y G D VM I F R S I D Q H VV M P T I T LR F NI Y T D PRT P VQ VEPGLRA INDPGP-DD PVFI TTNFA LT YYTVES D - - - - - - - - - - G I S D P V S A A Y WE T A M AA I F T I R Y G D IM I L H S L E P Y A TL P E V H L A E TI Y T D PRT P VA VDSKMYK VGEPDE-N SPVLF TTNFA LT YYTVES D - - - - - - - - - - G I A D P V S A S Y WE T V M AS V F T I R Y G D IM I L H S L E P Y A TL P E V H L A E TI Y T D PRT P VS VDGGMYK VGSPTA-D SPVLF TTNFA LT YYTVES D - - - - - - - - - - - H D D E V S A S Y WE T V V AS I F T I K Y G D IM I M H S I E P Y AL M P Q L H IR D TM Y T D PRK P VT VDPAVYE IGSPTA-D SPLLV TTNFA LT YYTVES D - - - - - - - - - - P E L S D D L N H W QE A Y L T S M L I TR Y A D LL I M H S I E G W AL L P Q L I WR F GI Y T D PRK P VS VDAGVRTFGNPDK-T SPVLI TSNYA LT FFTVES D L K N E C P I S G F F E D K E V V A K M FE A T I AN T L M N R Y A D AL I M H G M D I W EL M P V L T LR Q CI Y T D PRK PQA VEPGLYP IGNPDE-N SPVIL TTNFS LT FYTVTG D - - - - - - - - - - - E D D P V K A A R RE S Y L AS A C V L R Y A D IL I M D T V E P W AL L P V L T QR Q CV Y T D PRE PQE VEPGLYR IGDPDE-N SPVLV TTNFT LT YHCVAG D - - - - - - - - - - P E E D E I L T K M KE A T T AA A L M N R Y A D VM I L H G V D I W EM M P V L T LR Q SL Y T D PRK PQS VEAKIYEFGTVDE-N SPVIM TTNFA LT YYTVAG D - - - - - - - - - - - V S D K I E A N I RE A T V AA T L M N R Y A D IL I L A G T E I W EI M P V L T LR Q GL Y T D PRK PQA VDPGVYEFGDVDE-N SPVIL TTNFS LT YYTVEG D



1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

2 2 2 2 2 2 2 71 60 86 76 77 78 54 50 61 44 46

69 69 63 63 63 63 64 159 146 181 166 165 168 150 129 139 120 120

Mo.thermo D.hafniens C.hydrogen A.metallir C.difficil D.ethenog S.fumaroxi ANME1 ANME2 A.fulgidus Ms.thermo Ms.acetiv M.burtonii M.hutchins Mc.jann M.kandleri Mc.maripal M.thermaut

Mo.thermo D .h af ni en s C.hydrogen A .m et al li r C .d if fi ci l D .e th en og S .f um ar ox i A NM E1 A NM E2 A .f ul gi du s M s. th er mo M s. ac et iv M .b ur to ni i M .h ut ch in s M c. ja nn M .k an dl er i M c. ma ri pa l M .t he rm au t

M o. th er mo D .h af ni en s C .h yd ro ge n A .m et al li r C .d if fi ci l D .e th en og S.fumaroxi ANME1 ANME2 A.fulgidus Ms.thermo M s. ac et iv M .b ur to ni i M .h ut ch in s Mc.jann M .k an dl er i Mc.maripal M .t he rm au t

DVL KDP FTDV INE PGR WAQ KCVAE YG ADLIY LK LDGA D PEGANHSVDQCVATVKEVLQ AVGVPLVV VGCGDVE KDHEV LEAVAEA A AGENLLL G NA EQEDIV KSQ YGDV IND PVA WAKKCVEE YN ADMIY LK LIGANPEGENRSPEECA KVVKDVLA AVGVPLIV AGCEDPEKDNEV LAAVAEA T SGENLLI G IA EQDDIL V E PFKDV IND PVA WAKKCVE- YG ADIVA LR LVSAHPDGQNRSGAELA EVCKAVAD AIDVPLMI IGCGVEE KDAEIFPVI GEA L SGRNCLL S SA TKDASL KEL YKDV AND PAA WAKFVEEKLEPDFIC LRFEGSD PNGLDKSPEECA EVAKAVVE AISIPLVI AGTQNHDKDAKI F E K I T A A V DGHNCLF L AA TEEDSY KEL YKDV ANC PVE WAKYVEANTQADFIC LKFDGSD PNGLDKSVDECA DVAKAVIE AIKLPLVV AGSGNHE KDGKLFEKL AQ T L DGHNCLF M SA VEDEDV L K PFGDA VNN PVS WAKKCVDT YG AELIC LQ LESTD PNGLDRSAEESA KIVREVAD AVDVPLIV WGTANHE KDTEV LRKASEACPDKKLIL G P V E E G EAA KAP FLDV LADSAAWARKCVDQ YG AEMIV VQTKSA D PNGDNKPAEEVA EVVSKVIN AVDVPVVI WGVANHE KDTEV MRLTAEKCSDKRLAL S P V E E G KA V RGF YDDV MED PAE WAKKNVRE FG ADMVT IH LISTD PTIKDTSAKEAA KTVENVLQ AVKVPIVI GGSGNPD KDPVV LEAAAEA A AGERCLI A SA N L NM KA V RGH YEDV MED PGE WAKKNVRE FG ADMVT IH LISTD PTIKDTSAKEAA KTVEEVLQ AVKVPLVI GGSGNPD KDPGV LEAAAEA A EGERCLI A SA N L NM KAV RMH YEDV LDN PAE WARKCVKK FG ADLVT LH LISTD PLLDDTPASEAVKVLEDVLQ AVKCPIIV GGSGNKE KDPEV LEKAAEV A EGERIML A SA T L DM KAV KEN YDDV MDN PGE WAKKNVEK FN ADMIT IH LISTD PLIKDTSPKDAA KTVEEVLQ AVDVPIAI GGSGNPQ KDPLV LAKAAEV A EGERCLL A SA SLNL KAV KEN YDEV MDS PGE WAKKNVEK FN ADMIT IH LISTD PLIKDTPAKEAA KTVEEVLQ AVDVPIAI GGSGNPQ KDPEV LARAAEV S EGERCLL A SA SLNL KSV KEN YGDV IND PAE WAKKVVKD FN ADMVT IH LISTD PLINDTPAREAA KVVEDVLQ AVKVPIVI GGSGNPK KDPEV LERAAEV A EGERVLL A SA SLNL KAL KNHLTEV IDD PAA WAKMNVEQ FG ADMVT VH LLSTD PLIQNKSPKEASKTVEEVLQ AVDVPLII GGCGDPK KDAETFTEIAA M A EGERLLL N S V T S DM KPI RQF FQDV MED PCE WAKKCVKE FG ADMIT IHHISTD PKIKDKSPKEAA KLMEDLLQ AVDVPFVI GGSGNPQ KDPLV LEACAEV A EGDRCLL A SA NLEL GPI REELGDV IED PVD WARTVVKR YGV DIVT VH LVSTS PKLHDAPVEEAMETLEDILD AVKVPIIV GGSGDPE KDVEV FVKAAEV C EGERVML S S I N E DM RPI KEN FEDV MHS PGE WAQ KAVKE FG ADMIT LH LIGTD PKVKDKSLKDAA KDLEEVLQ AVKVPLVI GGSGNPK KDPLV LEMAAEV A DGERCLL A SA NLDL RPI REH FSDV MED PGD WARKAVKE YG ANMVT IH LIGTG PKVMDKSPREAA KDIEEVLQ AVDVPLVI GGSGDPE KDPLV LEKAAEA A EGERCLL A SA NLDL

------------------------------AVQILR-DRSRAAV Q KVVLG ATKDQGGTRSHTIVVG GDAA L PF H HFEGEIVNRPVI GMEVQ DI V P - - D W P ------------------------------AVALVK-ERYQSKV GEVVIG ATAAEGGTRTSVVKVG GDST L PF L HFEGKVENRAAI ALEVT DV P P - - A WN ------------------------------AVEVLK-EKWNSKV VEVTLG -------TGDKTVTLG GDST L PF L TFEGEM P NPPR F ALEVF DT P P T - D W P ------------------------------P FKMSV-QDFKGKI NEVEIG K-------GEKAIKIG GQKATALYHFDNDAGNAPKI GIEIT DI Y P E - E W I ------------------------------A FKMST-QKYSGKI SEVEVG I-------GEKAIKLG GENV L PF Y SFDGEVGNSPKI GIQIS DV Y P E - S W T ------------------------------A FEIPK-INYNGRI KEITLG E-------GPKAVTVG GETS M PF Y LFEGEM P RKPKI AMEIQ DT P P E - D W P ------------------------------A FQPAK-QVYTGAI REVTLG S-------GDKAVTVG GKKVFPF H SFEGTI P HPPRV AMEVW DK D P S A E W A I------QMLQPVQHVEEK-KV-AVLED-AK FEVAK-AAWTGKI EEVTIG ATSADGGTRESTVTIG GEKS L PF Y NFDHQM P YPPVI SVDCFDM A I - - P L A -------VQKVPIPTTPAE-EL--GLIR-TE FKVAK-TEWKGQI EEVTIG ATSADGGTREATVTIG GEKA L PF Y NFDHQM P HPPVI SVDCFDM P I - - T L A QLAPAAPAAPAEVPAVEEL-KIPAELVR-AK FEPYK-EEYPGKI EEVVLG ATKADGGTREYTVTLG GERS L AF Y TFDAPQ P HLPAI AIDVF DR R P - - M L A -----PAAAPAAVSPALAAPKL-KDLIP-AK FEFSNIEEWTTQV EEVPIG NTSADGGSRGKRILLG GEKA L PF F - PDAPM P NRNQV TIDVF DM R L - - G L A -------PAAPAVSPALAAPKL-KDLIP-AK FDVANIAEWATEI QEVPIG NTSADGGSRGKRVMLG GEKA L PF Y -FDAPM P NRNQV TIDVF DM R I - - G L A ----SMGGVPQQAPVAGAS-RIP-ELLP-AK FDVSKMSEWATPI QEVTLG ATSADGGSRKSTVTLG GENA L PY Y -FDAEM P HRNYV TMDVF DM P I - - S M A FPAPVAAAAKQMTPPVKRLPRIPDDLIS-EK FLMEP-DVFPAQI REITLG ATRAEGGTRGSTITVG GSTS I PFTHPQSPPP HPPVI SLDVF DT R V - - P M P LAEEGI--------------EI-KDVPE-LD WEPPV-EKYPGYI REVQFG KPKSEGG-RGKVVKIG GQRA L - - Y RFEEPQ P NPPVV T FDIF DI P M P - G L P EVEEKER-------------E--VEVPE--- YEPPV-EEYEGCV AEVQIG ATRSDGGSRDRVVVLG GERA--Y F PFEEPR P NPPVV T FDVF DT P D V - G I P -VVEKQI-------------EAKKEFESQIE FKYPV-KDYAGKV NEVKLG ------GKNRKTVKLG GQTS L - - Y RFEEPQ P NAPTV T FDVF DI P M N - G L P -TVEVR--------------EAVEALPAE-E FNPPI-RSYPGEV A QVKLG E-----GTR-KSVYLG GQKA L - - Y RFEEPQ P NPPVV T FDVF DI P M P - G L P

M------------------------------------------------------------------------------------------- - - - - - - - M------------------------------------------------------------------------------------------- - - - - - - - M------------------------------------------------------------------------------------------- - - - - - - - M------------------------------------------------------------------------------------------- - - - - - - - M------------------------------------------------------------------------------------------- - - - - - - - M------------------------------------------------------------------------------------------- - - - - - - - M------------------------------------------------------------------------------------------- - - - - - - - M--RKKITFGDLASILE-------NYDIVELEDVKIE-GDLEVELSQSG-AIPP-PLI-Q-VLQSMLNEVDKQ-------VEAT-----LT- - - - T R K S L M--EKEITLADIAKILE-------KYEIVELEDVKIK-GDVVLELAAGGVALPPSIV--Q-ALQSIIS------------------------ - - - A G A G M---KKFTLEEFLDLLK-------KYNVEEIEGVKIE-GDLEIEFEGSS--IDLSAIAPLYSMLSEFGNFLYHANMALGYLQRLSAALGIPT P - - F A A Q P M--AKKMKLSEITNMFA-------GMNVEALEGVTIE-GDIEIDLGGLGGF-DPMLA--A-ALGQESA-VLAQ------HFARIAGMFGYPV S - - - V G A M--AKKMKLSDITNMFA-------GMDVEALEGVTIE-GDIEIDLGGLGGGFDPMLA--A-ALGQESA-VLAQ------HFARLAGMFGYPV G - - - I G A M--SNKMKLSGLTDILK-------DLDVQSLEGVTIE-GDIELDISGGGGL-NPAIA--Y-ALGHEAA-QISM------HVANIARMLGYPV D Q L F A A - MTDQEKPDISNLIRLLGPGLGELLSGRQIELEHVELELGELELWIPVGG------------------------------------------- - - - - I S Q G M------DLNTLIKIIE-------KVGRIEIEDIKITA-D-ELIIN-----IPSAPPI---VIPQTPSIKE--------------------- - - - - - - - K M--AEDKGVKNMLEHLVKNL-LVEDVEEIELRNVTIELDELELDLKTVA---------------QTLPVEI--------------------- - W E R I L K P M-----DTMSQILKLLE-------KTDSIEINEFRMDV-D-ELELT-----IM--PTL-Q-NVVQK-------------------------- - - - - - - - M---LMDKLTELLKLLQ-------NTESIEINEFRMDV-E-ELELY-----LMPA-V--Q-QAIQK-------------------------- - - - - - - - -


Appendix 5. Alignment of δ Subunit Protein Sequences

Met. Ions Life Sci. 2, 357–416 (2007)

1 68 1 68 16 1 1 62 1 62 1 62 1 63 2 59 2 46 2 81 2 66 2 65 2 68 2 50 2 29 23 9 2 20 2 20

2 61 2 61 2 54 2 57 2 57 25 6 2 57 351 338 3 74 35 9 35 8 3 61 3 45 32 1 3 31 3 12 3 12

M o. th er mo D. h afn i ens C.h y dro g en A. m eta l lir C .d if fi ci l D .e th en og S. f uma r oxi AN M E1 AN M E2 A. f ulg i dus Ms . the r mo Ms . ace t iv M. b urt o nii M. h utc h ins Mc . jan n M.k a ndl e ri Mc . mar i pal M. t her m aut

Mo . the r mo D .ha f nie n s C .hy d rog e n A .me t all i r C .di f fic i l D .et h eno g S .fu m aro x i ANME1 ANME2 A .fu l gid u s M s.t h erm o M s.a c eti v M .bu r ton i i M .hu t chi n s M c.j a nn M .ka n dle r i M c.m a rip a l M .th e rma u t

Met. Ions Life Sci. 2, 357–416 (2007)

KEASA---PVSEYPGW G KE TER GI L WEAV TA T A LLQA G AH IL LM RHP EAV A RV KENIDQLMVSNAY-------------------- KEANA---AEEEFPGW G DL NDR AI L WEAL TA A G LLQV G AS IL LM RNP AAV K LVQQNIADLMEMNA--------------------- KEAKD---P--EVAEW G DY ALR A I H WETV TT V ALIQA G GH LF VM RHP KSL A EV KEHLKRILK------------------------ KESIA---EEEDAPEW G SR EER A I S IEISTA T S IIAV G ADAV IV RHP KSL E TL KNLTSQLA------------------------- KEAIA---PIEDEPDW G CP EERT I A MEVSTA A S VLVGG SNAV IL RHP KSI E TI KELVNALA------------------------- KEAKI---PEAESPEM G NT EKR AI M IEAM SA A I LLIAG AD IL VI RHP ESMRLAGELIDELS------------------------- KEANL---SAEDAPAL G DP ARR GI L MEAV TA V G LLLAG AD IV VM RHP DAI K LV KQFIAKMT------------------------- RESWMKGSPLAEDSDW G PA AYR GP L YEII TG L T LGIAG GD MF MM MHP KAAAAV KRVTQMLFGAD-T-DGAKEVNVNEWVTWEANK- REAWMKDSPIREDSDW G PA ANR GP L YEIM TG L T LGIAG GD MF MM MHP KAAAAV KRITQMLFGATGAADTGKKVKMEDWVTWGGV-- REAWMVDSPIEEDTPW G PR ELR GP I WEII TG L T LSLAG VD LF MM MHP VAV A VL KEVFNTLGGKVSG----GVADPGEWIFMEV--- RESWMVSSPLKEDSDW G PR EYR GP I WEII TG L S LAIAG ND LF MM MHP TSV A VL KHMTQTLFGSIEA----EPVDIANWIGAEV--- RESWMVSSPLKEDSDW G PR EYR GP I WEIV TG L S LAIAG ND LF MM MHP TSV A VL KQITQTLFGMIDT----EQVDVANWIGAEV--- RESWMKESPLKQDSDW G PR EYR GP I WEII TG L T LSLAG ND MF MM MHP TSV Q VL KEITQTLYGSIEA----EEIDITKWIGAEV--- REAWM-----DLGPAF G RN DLR GP L WETIGG I T LLLAG VD LF LM MHP LAV S TL RDVAGKLMHPAEI----KP-DMADWAALKI--- REAWM------NNPEW G PR EYRLP L WEITTG I T MMMCG VD LF MM LNP ISV K TL KEIGKTLTTKPGE--VKLNTNNYEWIVSP---- REAWM------KEESW G PR EYR GP L WEAV TA T T VALCG AD LL MM FHP WAV Q VVMEAMEYMAEGRVT----GDAYVTDVIA------ REAWM------KKDEW G PT KYR GP I WEVV TG L T MMLCG VD IF MM LHP LSV K TL KEIGTTLTYEY---KSKESQDISDWINKF---- REAWM------KKDEW G PT DYR GP L WEIV TG L T MMLSG VD IF MM LHP TSV R LL REIGETFTREY---MTAETPDLREWITELDYQE V

--- -NYK SL T A-A CM VH K HNIIAR S PLD I NI CKQL NIL INE-MN LPLDHI VI D PSIG G L G YG IE YS FSI ME RI R LGAL -QG DKML S MPV I C T V G Y EAW RA --- -NYK SI T A-A AM VH K HNLIAR S PLD I NI CKQL NIL ISE-MG LPLNRI VI D PMIG G L G YG IE YA YSI ME RA R LGSL -AN DKML S MPM I C T V G Y EA N R A -- -- NYK PI V A- TC MV HGHSVV A S A PLD I NL SKQL NIM IME-MN LAPNRI IM D PLIG A L G YG IE YS YSI IE RM R LGAL -TG DKIL A MPV V C F I G Q EAW KA --- -NYK TV G ASA GL AY K HKVGAE S SVD I NL AKQL NVL LDQ-LG VKNEDI TMHIGCS A V G YGFE YL IST VD RI R LAAL GQN DKTL Q MPI I T P V S F E TW TV --- -NYK GV G ASA GM AY A HKVGAE S SVD I NL AKQL NVL LTQ-LG VKGENI VMNVGCS A V G YGYE YV AST MD RI R LAAFGQN DKTL Q MPI I T P V A F E V G HV --- -DYK KI A A-QAM AY K HTIIAS S PID I NL AKQL NIL LGN-LG VPAEGL IM D PTVSS I G YG IE YS YSV IE RI R LAAL TQQ DERL Q YPL I C N I G R EAW KT --- -DYK QV G A-A CL AY K HVVVAS S PID V NL AKQL NIL LGN-LG VAGQDI VI D PTTG G L G YG LE YS YSV ME RI V Q AAL TQE DEKL Q QPI I A N V G N E VW KC - -- -DYE RI - AKA AQ DN G HVILSW T QLD I NA QKTL NRY LFK-LG VPREDI VI D PTTA A L G YG LD YA FTN IE RM R IAAL -KG DTDL A FPI S C G V - T NAW GA - -- -DYE RI - AKA AL NH G HVILSW T QLD I NA QKTL NRY LFK-LG VPREDI VI D PTTA A L G YG LD YA FTN IE RM R ISAL -KG DTDL A FPI S C G I - T NAW GA --- -DWE RI - ANA AK KY G HVVLSW T QMD I NN QKTL NRY LLKRVE MPRDSI VM D PTTA A L G YG LD YA FTN ME RI R ISGL -KG DTDL N FPI S S G T - T NAW GA ----DYA AI - AEA AL KY D HDVLSW T QLD M NA QKEL NRK LMKQCN VPRDRI IM D PTTA A L G YG LD YA YTN ME RI R LAAL -MG DDEL T FPM S S G T - T NAW GA ----DYA AI - AEA AL KY D HDVLSW T QLD M NA QKEL NRK LMKQCN VPRDRI IM D PTTA A L G YG LD YA YTN ME RI R LAAL -MG DDEL T FPM S S G T - T NAW GA --- -DYE RV - AKA AT DH G HAVLSW T QLE I NA QKEL NRK LMKQCN VPRDSI VM D PTTA A L G YG LD YA YTN ME RI R LAGL -MG DDEL T FPM S S G T - T NAW GA ADA KT LE PV - C KA AD TH G HCLLGF T GLD L NS AKEL NRR IY-QY-FPPERL LM DLTTV A L G YG LE YS FSIHE RA R MAAL -MG DPEL Q HP T I S A C - T NAW SA ----DYK KI - V DA AM KY D HNVLAW S IMDP NM ARDL NRK LVE-AG LDPNRI VM D PTTC A L G YG IE FSINA MV RL R LNGL -KG DELV N MPM S S G T - T NA I G A -- -- DFE RV - V EA AK EH G HVVL T F A PVD V NL MKSL NKK VLN-RG LSKEDV VM D PTTC A L G YG IE YT I D VMT RI R LAAL -KG DEHL Q MPI S S G S - T NAW AA --- -DYQ KI - AQA AV KH D HAVLSWAISD I NM QKVL NKA LMK-EG LTANDI VM D PTTC A L G YG IE FSIDV MT RT R LSAL -KG DEIL Q MPM S S G T - T NAW GS --- -DYR KV - ARA AL DH N HAVLSWAITD V NM QKTL NRY LLK-EG LKREDI VM D PTTC A L G YG IE FSIDV IT RT R LAAL -KG DSDL Q MPM S S G T - T NAW GS

Appendix 5. (Continued) 408

Subunit

sp冟Q8TXF4 ref冟ZP_00562509.1 sp冟Q8TRZ6 sp冟Q8TJC4

Methanococcoides burtonii Methanosarcina acetivorans

sp冟Q57616 sp冟Q57620 ref冟NP_988103.1 ref冟ZP_00869638.1

sp冟O27745

sp冟O29868

gi冟82617264 gb冟AAU83256.1

α

sp冟Q57617 sp冟Q58138 ref冟NP_988105.1 ref冟ZP_00869636.1 ref冟ZP_00868193.1 sp冟Q8TXX3 sp冟Q8TXF7 genome sp冟Q8TRZ4 sp冟Q8TJC6 sp冟Q8TKW2 sp冟Q8TR73 sp冟Q8THW2

gb冟AAU84344.1 sp冟O28429 sp冟O30274 sp冟O29165 sp冟O27743

gb冟AAU83593.1

β

γ

ref冟ZP_00562513.1 gb冟AAM04441.1 sp冟Q8TH44

sp冟Q8TXF1

ref冟NP_988100.1 ref冟ZP_00869640.1

sp冟Q57576

sp冟O27748

sp冟O29871

gb冟AAU82436.1 gb冟AAU83253.1 gb冟AAU82655.1

Subunits (GenBank or Swiss-Prot accession numbers)

Methanopyrus kandleri

Methanococcus maripaludis Methanospirillum hungatei

Methanothermobacter thermautotrophicus Methanococcales Methanocaldococcus jannaschii

ANME clones ANME Fosmid GZfos27B6 ANME Fosmid GZfos18H11 ANME Fosmid GZFos9D8 Archaeoglobus fulgidus

Euryarchaea

Organisms

Appendix 6. Protein Sequence Identifiers and Accession Numbers

(Continued)

ref冟ZP_00562512.1 sp冟Q8TRZ8 sp冟Q8TJC2

sp冟Q8TXF2

ref冟NP_988101.1 ref冟ZP_00869639.1

sp冟Q57577

sp冟O27747

sp冟O29870

gb冟AAU83254.1 gb冟AAU82656.1

δ


CD0728 ref冟ZP_00558638.1

sp冟P27988

ref冟ZP_00665908.1

Clostridium difficile(Sanger Institute)

Desulfitobacterium hafniense

Moorella thermoacetica

Rhodospirillum rubrum Geobacter sulfurreducens Syntrophobacter fumaroxidans ref冟YP_307738.1

ref冟YP_360060.1

Carboxydothermus hydrogenoformans

Dehalococcoides ethenogenes 195 Chlorobium phaeobacteroides

ref冟ZP_00800477.1

Alkaliphilus metalliredigenes

Bacteria

sp冟Q9V2Z4

Methanosarcina thermophila

α sp冟Q8PZ11 sp冟Q8PV85

Subunit

ref冟ZP_00528138.1

ref冟ZP_00801249.1 ref冟ZP_00800465.1 sp冟Q9F8A8 ref冟YP_359590.1 ref冟YP_358906.1 CD0174 CD0716 ref冟ZP_00558637.1 ref冟ZP_00097361.1 ref冟ZP_00099357.1 ref冟ZP_00560129.1 sp冟P27989 ref冟ZP_00576900.1 sp冟P31896 ref冟NP_953147.1 ref冟ZP_00664754.1 ref冟ZP_00665907.1

sp冟Q8Q0L5 sp冟Q8PUN1 sp冟Q49161 sp冟Q49163 sp冟Q9C4Z4

β

γ

ref冟YP_181407.1

ref冟ZP_00665909.1

sp冟Q07340

ref冟ZP_00558639.1

CD0726

ref冟YP_360061.1

ref冟ZP_00800475.1

sp冟Q9C4Z0

sp冟Q8PZ09 sp冟Q8PV87

Subunits (GenBank or Swiss-Prot accession numbers)

Methanosarcina mazei

Organisms


ref冟YP_181409.1

ref冟ZP_00665906.1

sp冟Q07341

ref冟ZP_00558642.1

CD0725

ref冟YP_360064.1

ref冟ZP_00800474.1

sp冟Q9C4Z1

sp冟Q8PRQ5 Gb冟NP_634109.1

δ



411

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G. Wächtershäuser, in The Molecular Origins of Life: Assembling Pieces of the Puzzle (A. Brack, ed.), Cambridge University Press, Cambridge, 1998, pp 206– 218. P. A. Lindahl, Orig. Life Evol. Biosph., 34, 371–389 (2004). M. J. Russell and W. Martin, Trends Biochem. Sci., 29, 358–363 (2004). C. Huber and G. Wächtershäuser, Science, 276, 245–247 (1997). F. Ronquist and J. P. Huelsenbeck, Bioinformatics, 19, 1572–1574 (2003). J. Felsenstein, PHYLIP (phylogeny inference package), 3.65; Department of Genetics, University of Washington: Seattle, 2005. R. E. Duderstadt, C. R. Staples, P. S. Brereton, M. W. Adams, and M. K. Johnson, Biochemistry, 38, 10585–10593 (1999).

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10 Nickel Superoxide Dismutase Peter A. Bryngelson and Michael J. Maroney Department of Chemistry, 701 Lederle Graduate Research Tower, University of Massachusetts, 710 North Pleasant Street, Amherst, MA 01003-9336 <[email protected]>

1.

2.

3.

4.

5.

INTRODUCTION 1.1. Superoxide Dismutase Enzymes 1.2. Nickel Redox Chemistry MOLECULAR BIOLOGY 2.1. Streptomyces sp. 2.2. Expression, Processing, and Metallocenter Assembly STRUCTURAL BIOLOGY 3.1. X-ray Absorption Spectroscopy 3.2. Crystal Structures MECHANISM 4.1. Spectroscopy 4.2. Kinetics 4.3. Model Studies CONCLUSIONS ACKNOWLEDGMENTS ABBREVIATIONS AND DEFINITIONS REFERENCES



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INTRODUCTION Superoxide Dismutase Enzymes

Superoxide dismutases (SODs) are oxidoreductases that catalyze the conversion of superoxide to peroxide and molecular oxygen at physiological pH and at rates near the diffusion limit (kcat ⬃ 109 M1s1). As such, SODs are important components of systems that protect biological molecules from oxidative damage by superoxide or reactive oxygen species (ROS) generated from superoxide, and specific deficiencies have been associated with defective SODs [1,2]. Superoxide radical has been linked to the aging process and to various disorders and diseases ranging in severity from inflammatory to neurodegenerative [3–6]. The latter includes familial amyotrophic lateral sclerosis (fALS), which has been linked to point mutations in CuZnSOD [7], and Parkinson’s disease [8], where increased expression of CuZnSOD in primary dopaminergic neurons increases their resistance to the neurotoxin 6-hydroxydopamine. Superoxide-mediated damage to mitochondrial function is thought to contribute to organ failure caused by the transient disruption of the blood supply in ischemic diseases, especially heart disease [9], stroke [10], intestinal ischemia [11], and organ preservation for transplantation [12]. Alcohol-induced liver damage has been shown to require superoxide, and overexpression of MnSOD moderates this effect [13]. SODs have also been shown to be important virulence factors in bacterial pathogens of humans and domesticated animals, and are therefore potential targets for drug therapies [14–16]. Nonetheless, there is no known advantage conferred on microorganisms that possess NiSOD, rather than the more common Fe-, Mn- or CuZn-containing enzymes. All known SODs are metalloenzymes that utilize metal-centered one-electron redox chemistry to catalyze the disproportionation of two molecules of superoxide to a molecule of hydrogen peroxide and molecular oxygen [1,17]. The mechanism by which SODs achieve catalysis involves electron transfer to and from the redox-active metal site coupled with rapid proton transfers (PCET) [18], and proceeds via a ping-pong mechanism wherein the metal is first reduced and then reoxidized by superoxide (Equations 1–3). M( n +1) O•2 → M n O2

(1)

M n + O•2 2 H → M( n +1) H 2 O2

(2)

2 O• 2 2 H → O2 H 2 O2

(3)

Many well-characterized examples of SODs with active sites containing Mn or Fe, Mn/FeSOD, or a bimetallic CuZn, CuZnSODs, are known [1,17,19–22]. FeSODs are found in prokaryotes and in plants, while MnSODs are found in Met. Ions Life Sci. 2, 417–444 (2007)

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prokaryotes and the mitochondria of higher organisms, including humans. The CuZnSODs are commonly found in the cytosol of eukaryotic cells and in the perioxisomes, but have also been found in the periplasmic space in some bacteria. Recently, a third class of SOD with Ni as the active site metal has been characterized from microorganisms, notably Streptomyces sp. [19–22]. The NiSODs have many similarities with the Mn/Fe and CuZn SODs, but form a distinct group based on the metal content and ligand environment of the nickel center, the protein structure, and several aspects of the reaction mechanism. A requirement for the active-site metal is a one-electron redox potential that lies between the potentials for the oxidation and reduction of O 2 . The O2 /O2 couple has a reported reduction potential of 160 mV and the O2 /H2O2 redox couple is 870 mV [23]. The midpoint potential for Equations (1) and (2) is roughly 360 mV in aqueous solution at pH 7. Mn, Fe, and CuZn SODs have measured redox potentials that essentially center on 300 mV with a range of ± 100 mV [24–28]. The Cu(II/I), Mn(III/II), and Fe(III/II) redox couples in the respective SODs supply the requisite one-electron redox center. Many aquated metal ions, including Cu2, Mn2, and Fe2, are capable of SOD activity, but Ni2 is not among them [1]. Because the Ni(III/II) couple lies at over 1V and the Ni(II/I) couple is lower than 1 V in aqueous solution, Ni has only one common oxidation state in water, Ni(II). Nickel is therefore an unlikely metal center for a SOD. Nonetheless, a similar redox potential for NiSOD (286 mV) has recently been measured [29]. Since oxidized NiSOD exhibits an S 1/2 EPR spectrum appropriate for a Ni(III) center [30], catalysis relies on a protein environment that is capable of lowering the Ni(III/II) couple by 1 V. The primary amino acid sequence of NiSOD shares no sequence or structural homology with Mn/FeSODs or CuZnSODs [31]. Mn/FeSODs and CuZnSODs share common active site features that include metal ligand environments composed of N- and O-donors derived from amino acids (histidine and aspartate) as well as an aqua/hydroxo ligand [32–34]. The amino acid sequences of Mn and Fe SODs are highly similar, although Mn-substituted FeSODs (and vice versa) are generally inactive [25]. Even though their active sites are virtually identical, hydrogen bonding networks involving second sphere residues play an important role in optimizing the one-electron redox potential of the active site [25]. In contrast, the nickel active site in NiSOD incorporates cysteine thiolate S-donors, three different types of nitrogen donors, and lacks an aqua/hydroxo ligand. The strategy of using cysteine-thiolate ligands to access the higher oxidation state (in this case the Ni(III/II) couple), is a very common one in other nickel-redox enzymes [35], but is unprecedented in SODs, and unexpected because of the facile oxidation of thiolates, including metal thiolate ligands, with H2O2 and O2 [36,37]. In addition to the metal redox chemistry, two protons are required for the production of H2O2, the second of which assists product release. Thus, the enzyme catalyzes a one-electron, two-proton process. This is accomplished in Mn/FeSODs by a series of proton-coupled electron transfer reactions, where uptake of H accompanies reduction of the enzyme by superoxide, and reduction of O2 is also Met. Ions Life Sci. 2, 417–444 (2007)

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Figure 1. The dependence of kcat on pH for NiSOD (solid circles), MnSOD (squares) and CuZnSOD (triangles). Adapted from [30] by permission of the American Chemical Society.

accompanied by uptake of H [18,38]. Thus, Equations (1) and (2) can be rewritten to reflect the PCET reaction mechanism: E ox O•2 H → E red (H +) O2

(4)

E red (H + ) O•2 H → E ox H 2 O2

(5)

By supplying a proton for the reduction of O 2 , the enzyme is able to couple a favorable reaction (oxidation of O to O ) to drive an unfavorable one (reduction 2 2 of O ). Uptake of a proton upon reduction of the enzyme has been demonstrated 2 for Mn/FeSODs and CuZnSODs [18,39,40], but not for NiSOD, which is assumed to function in an analogous manner. Figure 1 compares the pH dependence of kcat for various SODs. This graph shows that, over a wide range of pH values, the enzyme catalyzes the dismutation of O2 at about 109 M1s1. At pH 4.8 (the pKa of HO2) the uncatalyzed rate is at a maximum (HO2 7 1 1 O 2 H → O2 H2O2 ; k 9.7 10 M s ) [1]. At intermediate pH values (pH 5–9.5) the catalytic rate constants are independent of pH, indicating that, although these protons originate from bulk solvent, the source of protons supporting the reaction is within the protein. A model where protons are transferred via a H-bonding network to the OH ligand in the oxidized Mn/FeSOD enzymes, with Tyr34 extracting a proton from nearby water, has been advanced, based on theory, and has considerable support from spectroscopic studies [38]. Despite the aqua ligand bound to Cu in CuZnSODs, the proton coupled to the reduction of the Cu(II) site is bound to the His63 imidazolate that bridges between the Cu and Zn ions and dissociates upon reduction and protonation [38,41]. The groups whose pKa values are responsible for loss of activity at high pH, differ with redox state and metal site. For FeSOD, binding an additional OH to Met. Ions Life Sci. 2, 417–444 (2007)


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Fe(III) in the oxidized enzyme, or deprotonating Tyr34 in the reduced enzyme account for the loss of activity [42]. MnSOD does not appear to bind additional hydroxide. Deprotonation of Tyr34 in the oxidized enzyme, and deprotonation of this residue coupled with partial deprotonation of the axial aqua ligand in the reduced enzyme may be the functional change in this system [42].

1.2. Nickel Redox Chemistry In order to access the Ni(III/II) redox couple in aqueous solutions Ni requires a ligand set other than water or neutral O- or N-donors (Table 1) [43]. The calculated value for the Ni(III/II) redox couple of aqueous nickel ions is 2.29 V (versus NHE), rendering it unattainable in water [44]. Nitrogen ligation can support a one-electron oxidation of Ni(II), and can vary the potential over a wide range (0.79 V to 1.72 V). Among these complexes are several Ni(II) peptide complexes that feature deprotonated amidate ligation, some of which can be oxidized by O2 [50]. The lower potentials associated with the amidates benefit from the negative charge associated with the ligands. However, the Ni(III/II) redox potentials supported by N-donor ligands are generally beyond what can be accessed by Table 1. Ni(III/II) reduction potentials of selected complexes. Given are the reduction potentials for a variety of nickel nitrogen-ligated complexes in aqueous solutions showing a wide range of reduction potentials for the Ni(III/II) couple. Sulfur ligation in NiFe hydrogenase is believed to lower the reduction potential substantially. Complex Niaq Ni(bipy)3 Ni([13]aneN4) Ni([14]aneN4) Ni[H3 (gly) 4] Ni-1* Desulfovibrio gigas hydrogenase [Ni(ema)](Et4N) 2 Desulfovibrio gigas hydrogenase

Coordination NiO6 NiN6 NiN4 NiN4 NiN4 NiS2N2 NiS4O NiS2N2 NiS4

E1/2 versus NHE (V) 2.29a 1.72b 1.12b 0.92b 0.79b 0.30 c 0.150 b 0.30 c 0.390 b

Reference [44] [45] [46] [46] [47] [48] [49] [48] [49]

a

Calculated. Measured in aqueous solution. c Measured in acetonitrile. Ni-1* (N,N-bis-2-mercapto-2-methylpropane-N,N-diazacyclooctane) nickel(II) Ni([13]aneN4) (1,4,7,10-tetraazacyclotridecane) nickel(II) Ni([14]aneN4) (1,4,8,11-tetraazacyclotetradecane) nickel(II) Ni(bipy)3 tris-(2,2-bipyridine) nickel(II) [Ni(ema)](Et4N)2 bis-tetraethylammonium [(N,N-ethylenebis-2-mercaptoacetamide) nickel(II)] Ni[H3 (gly)4] sodium [(tetraglycine)nickel(II)] b

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biological redox agents (0.42 to 0.8 V, roughly defined by the H2 /H and the O2 /H2O couples at pH 7) and too high to support efficient turnover of the SOD reaction [43]. Further lowering of the Ni(III/II) redox potential (even to negative values compared with NHE) can be achieved using thiolate (or sulfide) donors (Table 1), as has been illustrated in a number of NiN2S2 complexes [51], provided that oxidation of the ligands can be avoided (Equation 6). 2 Ni(III) 2 RS → 2 Ni(II) RS-SR

(6)

The usual strategies to avoid disulfide formation are to modify the electronic structure of the thiolate ligand by adding electron withdrawing groups (i.e., make S harder to oxidize), modify the Ni(III) center with the addition of good electron donors (i.e., make Ni(III) less oxidizing) or rely on sterics to encumber disulfide formation. In proteins, cysteine ligands can be positioned precisely and their interactions controlled by the protein environment. The electron-donor capabilities of cysteine thiolate ligands can be modified in proteins by H-bonding interactions and the hydrophobic/hydrophilic nature of the environment, the classic example being the range of potentials exhibited by Fe4S4 clusters in ferredoxins and HiPIP [52]. Additional anionic ligands on the Ni center also lower the redox potential and help favor metal-centered redox over S-centered redox. In addition to disulfide formation, oxidations of nickel thiolates by H2O2 and O2 (the products of the SOD reaction) are known to form a variety of S-oxygenates, including sulfenate, sulfinate and sulfonate complexes [36,37,53–55]. Based on synthetic model studies, the expected consequence of such oxidations in the enzyme would be to increase the redox potential of the Ni site to values that are no longer appropriate for SOD catalysis [53,56–58]. Sulfur K-edge X-ray absorption spectroscopy (XAS) provides a good assessment of the sulfur species in a sample [59,60], and shows no evidence to support the presence of S-oxygenates in the enzyme [61]. Similarly, no mass spectral data are available to show incorporation of oxygen in NiSOD. The apparent incompatibility of thiolate ligands with the production of H2O2 and O2 at the active site, and the manner in which NiSOD is able to protect the thiolates from oxidation is not understood, though protonation may be part of the answer. Such protection has also been proposed in order to explain the protection of an Fe-cysteinate ligand in superoxide reductase, an enzyme that catalyzes the reaction described by Equation (2) [62].

2.

MOLECULAR BIOLOGY

2.1. Streptomyces sp. First discovered in Streptomyces sp., sequences encoding putative NiSODs are now known in genomes of a larger variety of organisms [31,63]. While Streptomyces Met. Ions Life Sci. 2, 417–444 (2007)


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sp. contain sodN and sodF genes encoding the NiSOD and an FeSOD, respectively, addition of Ni to cultures of S. coelicolor results in a nearly 10-fold increase in the transcription of the sodN gene [64,65]. Along with up-regulation of sodN, the transcription of sodF is repressed. Addition of divalent transition metal salts (FeCl2, MnCl2, CuCl2, and ZnCl2) to growing cultures has no transcriptional or post-transcriptional effects on either sodN or sodF. Cobalt, the only other metal studied, has moderate effects. By regulating both superoxide dismutase genes, nickel appears to be the preferred metal and preferred SOD of Streptomyces sp. Micromonospora sp. [66], Nocardioides sp. [67], Frankia sp. [68], Janibacter sp. [69], Prochlorococcus sp. [70], Trichodesmium sp. [71], Crocosphaera sp. [72], Synechococcus sp. [70], Micromonospora sp., Microtetrasproa sp., Kitasatospora sp. [73], and Colwellia sp. [74] all contain genes for hypothetical proteins that have a high homology to sodN of Streptomyces sp. Expression of sodN from Prochlorococcus marinus shows that the organism encodes for an active NiSOD that can be expressed in Escherichia coli [75]. Micromonospora rosaria, Microtetrasproa glauca, and Kitasatospora griseola also have NiSODs which reproduce the EPR and UV/Vis spectra of the enzyme from Streptomyces sp. [73]. Amino acid sequences deduced from the nucleotide sequences for this highly homologous group of enzymes are compared in Figure 2.

2.2. Expression, Processing, and Metallocenter Assembly Common to all known sodN genes are N-terminal leader sequences of varying length that must be post-translationally processed prior to nickel incorporation and hexamer assembly. Expression of the full-length protein, encoded by sodN of S. coelicolor, in E. coli results in an inactive enzyme that does not incorporate nickel [76]. The peptidase responsible for post-translational removal of the 14 N-terminal amino acid residues in S. coelicolor is not present in E. coli. Some activity was originally seen in a truncated form of NiSOD (sodN∆), where residues 2–14 were removed; however, the production of active enzyme was attributed to modification of the protein by methionine aminopeptidase present in E. coli [64]. Several methods have been employed to produce active wild-type and mutant NiSODs. Native NiSOD can be isolated directly from S. seoulensis and recombinant wild-type and mutants can be produced in E. coli as well as in S. lividans TK24 ∆sodN (a NiSOD deletion mutant) or S. coelicolor A3. Production of NiSOD and mutants in Streptomyces sp. is the original method of obtaining suitable quantities of the enzyme for study. Spores are grown on solid media for 5 days at 28C, they are collected and used to inoculate liquid cultures incubated with shaking for an additional 2 days at 28C [77]. Addition of NiCl2 to the media enhances the expression of NiSOD nearly 10-fold [64,78]. The enzyme is processed by the internal molecular mechanisms of the Streptomyces sp. and the Met. Ions Life Sci. 2, 417–444 (2007)

Figure 2. Sequence alignments for NiSODs from a variety of organisms. Identical residues are shaded in black and highly similar residues are shaded in gray.

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425

metal is incorporated. Homogeneous NiSOD was obtained following purification, as described [31]. Heterologous expression of active NiSOD enzyme has been accomplished in two differing ways in E. coli that rely on either in vitro or in vivo post-translational processing followed by metal incorporation. The method for production via in vitro processing involves a sodN∆ construct lacking the nucleotides encoding for amino acid residues 1–14, that is inserted into the pET-30Xa-LIC vector [76]. The resulting His-tagged fusion protein expressed in E. coli is purified via IMAC. The fusion protein has the protease factor Xa amino acid recognition sequence, IEGR, prior to the start of the native processed sequence of NiSOD. Protease cleavage following the Arg residue produces the proper post-translationally modified 117 amino acid apo-NiSOD [76]. Nickel incorporation and hexamer formation are spontaneous in vitro after the reduction of the intramolecular disulfide bond between Cys2 and Cys6 with DTT [76]. An alternate method for producing active NiSOD in E. coli is the insertion of a similar sodN∆ construct into the pET-26b vector. The resulting fusion protein includes a pelB leader sequence that directs the protein to the periplasmic membrane in E. coli. Addition of NiCl2 at the time of induction coupled with the pelB peptidase present in E. coli produces active enzyme in vivo that is purified as previously described [79]. A number of point mutations of NiSOD have been reported, focusing on the role of the Ni ligands [76]. The physical properties associated with these mutations are summarized in Table 2, and mechanistic aspects are discussed in Table 2. Effects of mutation on the activity of S. seoulensis and S. coelicolor NiSODa. Mutation H1Q H1(A,C,D,L,N,W, or Y) D3A Y9A Y9F Y9L Y9Q Y9W E17A M28L R39A R47A

Activity in cell extracts [73] b

Activity by pulse radiolysis [29,76] c

None detected None detected 23% None detected 78% None detected None detected 45% None detected N/A None detected 89%

17% N/A 51% N/A 73% N/A N/A N/A N/A 93% N/A N/A

a

Activities of several NiSOD mutants relative to wild-type activity. Activity in cell extracts, using the cytochrome c assay, where S. seoulensis mutants are expressed in S. lividans Tk24 ∆sodN [73]. c Activity measured by pulse radiolysis of isolated mutants measured at pH 7.5 [29,76]. N/A not available b

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Section 4. As a result of mutagenesis studies it was concluded that the N-terminal His residue was a critical ligand that required N-terminal processing, while Met28 (one of three S-containing residues in the protein) was not required for the structure of the active site or for catalysis.

3.

STRUCTURAL BIOLOGY

3.1. X-ray Absorption Spectroscopy Studies of NiSOD using Ni K-edge XAS provided the first information regarding the active site structure (Figure 3) [30]. Using frozen solutions of native oxidized and dithionite-reduced enzyme, XANES analysis revealed the interconversion of five-coordinate pyramidal and four-coordinate planar Ni centers in the oxidized and reduced sites. The EXAFS data obtained provided the first information showing that the ligand environment of the nickel site in both the oxidized and reduced enzyme included multiple S-donor ligands, although the data were unable to discriminate between mononuclear and dinuclear structures. This result distinguished NiSOD from all other known SODs, none of which feature S-donor ligands, and provided information about how the redox potential of Ni center could be brought into a range appropriate for SOD catalysis. Fits of the EXAFS data also provided metric details of the sites. A single average Ni-S distance of 2.158(1) Å and single average Ni-N distance of 1.909(4) Å were reported for the oxidized sample, and a single average Ni-S distance of 2.154(4) Å and a shortened average Ni-N distance of 1.87(2) Å was found for the planar reduced site. Subsequently, XAS has been used to examine the structural consequences of point mutations of NiSOD (Table 2). Because the enzyme from Streptomyces sp. contains only three sulfur-containing amino acids (Cys2, Cys6 and Met28), it was correctly inferred from the EXAFS data that the Ni-binding site was at the N-terminus and involved ligation by Cys2 and Cys6 [30]. The low sulfur content of the enzyme also made it feasible to address the molecular and electronic structure of the site using S K-edge XAS (Figure 4) [61]. Model studies were used to show that different types of S-donors could be distinguished. The observation of pre-edge peaks in the S K-edge spectrum are indicative of the presence of covalently bound thiolates (and sulfide) ligands. These features depend upon electronic transitions involving promotion of the S 1selectron into vacancies in the Ni 3d-manifold. Since low-spin d7 Ni(III) has vacancies in both the dz2 and d x2y2 orbitals, two transitions are possible. In contrast, low-spin d8 Ni(II) has vacancies only in the d x2y2 orbital, thus only one transition is possible. Oxidized NiSOD includes two pre-edge features at 2469.7 and 2470.9 eV that confirm that one or more terminal thiolate ligands are bound to the Ni(III) center. Continued beam exposure photoreduces the Ni(III) to Ni(II), and the pre-edge peak at 2469.7 eV is lost. Retention of the peak at 2470.9 eV confirms Met. Ions Life Sci. 2, 417–444 (2007)


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Figure 3. XAS spectra for oxidized NiSOD (left) and reduced NiSOD (right): top, Ni K-edge XANES spectra; middle FT (k 2.0–14.0 Å1) EXAFS data (circles), fits (bold solid line) and differences between fit (solid line) and data; bottom FF( back transform window 1.1–2.6 Å, uncorrected for phase shifts.) EXAFS data (circles), fits (bold solid line) and differences between fit and data (solid line). Reproduced by permission of the American Chemical Society from [61].

that the Ni(II) center in the photoreduced sample also has at least one terminal cysteine ligand. NiSOD is also reduced by reaction with H2O2. Examination of the H2O2reduced wild-type NiSOD showed that the product of reduction with H2O2 (Figure 4) is distinct from that obtained by photoreduction. The S K-edge spectrum obtained from the H2O2-reduced sample exhibited only a broad edge and lacked any pre-edge features, indicating the absence of terminal thiolate Met. Ions Life Sci. 2, 417–444 (2007)

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Figure 4. Sulfur K-edge XAS spectra for NiSOD oxidized extrapolated (solid line), NiSOD photoreduced (dashed line), NiSOD peroxide reduced (dotted line), and free cysteine (dash–dot line) [61].

ligands. The S K-edge XANES spectrum of cysteine (and thioethers) show a similar broad featureless edge [76]. Thus, the data are consistent with protonation of the thiolate ligands upon reduction with H2O2. One representation of this reaction (Ni(III)SOD H2O2 → Ni(II)SOD ‘O 2 ’ 2 H ) is the reverse of the H2O2 producing reaction (Equation 2) and suggests that the thiolate S-donors are a possible source of protons for the reduction of O 2.

3.2.

Crystal Structures

High-resolution crystal structures are available for S. seoulensis [73] and S. coelicolor [79] NiSODs. These structures provide information about the protein structure, further refine the structure of the active site, and provide insights into catalysis by NiSOD. The structures of holo-NiSOD from these two sources are highly similar, as is expected from the 90% identity between the amino acid sequences of S. seoulensis and S. coelicolor NiSODs. The protein structure of NiSOD is distinct from either Mn/FeSODs, which are largely dimers, or dimers of dimers, that are dominated by α-helical domains with some β -sheet structure, or CuZnSODs, which are homodimeric or monomeric β -barrel proteins. NiSOD consists of a α6-hexamer with the mononuclear nickel active sites spaced 20 Å apart and arranged at the apices of a distorted octahedron. The hexameric quaternary structure is confirmed in solution by ultracentrifugation and ESI-MS [73]. The hexamer has a large void in the center of the enzyme, which is filled with water and co-crystallized ions. The presence of ions used in crystallization indicates that exchange of material is possible in solution, either through channels Met. Ions Life Sci. 2, 417–444 (2007)


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or flexibility in the quaternary structure. Individual subunits consist of four helix bundles that are stabilized by hydrophobic packing. The crystal structure of the S. seoulensis enzyme reveals that subunits corresponding to the apo-trimer are held together at one end by three His35-Gln37 H-bonds that lie on a three-fold axis. This interaction is not present in either the apo- or oxidized NiSOD of S. coelicolor, but a salt-bridge reported between Arg39 and Glu45 on alternating subunits holds this interface together [79]. Thus, the holoenzyme can be viewed as a dimer of trimers, where the three subunits of the trimers interdigitate. In addition to the interactions surrounding the threefold axis in the trimers, the nickel hook domain from one subunit of one trimer interacts with two of the subunits of the other trimer. These interactions include H-bonding between the His1 imidazole ligand of the nickel center and Glu17 from another subunit. Several other contacts hold the two trimers together, including hydrophobic, salt bridge and hydrogen bonding interactions. Near the active site pocket formed between two of the subunits on the opposing trimer there are several lysines that have been suggested to aid in electrostatic steering [79]. The crystal structures significantly advance the understanding of the active site structure by identifying the nature of two N-donor ligands (a backbone amidate derived from the N atom of Cys2 and the N-terminal amine) and by identifying the His1 imidazole ligand as the ligand lost upon reduction. Along with the thiolates provided by Cys2 and Cys6, the crystal structures show that all five ligands in the oxidized enzyme are derived from three residues in the N-terminus: His1, Cys2 and Cys6. The oxidized Ni(III) site is seen to be a five-coordinate pyramidal complex composed of three different N-donor ligands, (amine, amidate, and imidazole) and the two thiolate ligands. Upon reduction to Ni(II) the apical imidazole ligand, bound via the δ -N atom, is lost, forming a four-coordinate Ni(II) site, in agreement with XAS results. The first eight amino acids in the apo-enzyme crystal structure are disordered, but form a ‘Ni-hook’ domain upon binding nickel. Backbone amidate binding is an uncommon feature that has been observed in one other case involving a nickel metalloenzymes-CODH [80, 81].

4. 4.1.

MECHANISM Spectroscopy

Several spectroscopic techniques were used to elucidate electronic and structural information regarding the nickel site in NiSOD and its redox chemistry, including EPR, UV/Vis, MCD, and resonance Raman. The resting state of the enzyme exhibits a rhombic EPR spectrum with g-values of 2.306, 2.232, and 2.016 (Figure 6) [73]. This spectrum is typical of a low-spin d7 S 1/2 Ni(III) center with the unpaired spin in the dz2 orbital, and it shows the expected splitting upon incorporation of 61Ni (I 3/2) (Axyz 5, 5, and 30 G). Large N-hyperfine Met. Ions Life Sci. 2, 417–444 (2007)

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Figure 5. Crystal structures of apo-NiSOD (1T6I) and oxidized NiSOD (1T6U) from S. coelicolor: (top) Monomer from 1T6U showing four helix bundle and Ni bound at the N-terminus; (middle) Apo-NiSOD from 1T6I looking down the three-fold axis and perpendicular to the rotation axis. (Bottom-left) Hexamer formed by inter-digitation of two nickelated trimers; (bottom-right) Ni-hook region, amino acids 1-9 (HCDLPCGVY) with nickel bound.

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Figure 6. EPR spectra (upper) and simulated fits (lower) of NiSOD from S. seoulensis showing direct involvement of Ni with S and N ligands. (A) Native enzyme as-isolated; (B) 61Ni -enriched spectrum; (C) 15N-enriched spectrum; (D) 33S-enriched spectrum. Reproduced by permission of the National Academy of Sciences, U.S.A. from [73].

coupling is clearly observed for the g 2.016 signal, due to the 14 N (I 1) nucleus. The nitrogen interaction was confirmed by protein expression in 15 N (I 1/2) enriched media; the resulting EPR spectrum of NiSOD included a two-line hyperfine pattern in the g 2.016 signal, and a more detectable Nhyperfine interaction at other g-values (Axyz 22.7, 24.8 and 34.4 G) [73]. The N-donor ligand involved has been assigned to the apical His1 imidazole ligand on the basis of a theoretical model [82]. Hyperfine from 33S (I 3/2) (Axyz 3.6 G) was also observed [73]. EPR spectra from mutant NiSODs (Table 2) show two trends: either signals that are relatively unperturbed from the wild-type signal or no EPR. The unique EPR signal from oxidized NiSOD is lost upon reduction of the enzyme, indicating that the Ni(III/II) couple is utilized for SOD catalysis.

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The complex formed with the inhibitor N 3 (50% inhibition at 42 mM), shows an altered EPR spectrum with greater 14N coupling at gy [79]. However, this spectrum is unperturbed when 15N 3 is used, indicating that the perturbed spectrum results from structural or electronic changes that occur upon azide binding to the protein, and not from ligation of the nickel center. The UV/Vis spectrum of the oxidized wild-type NiSOD exhibits two absorption maxima associated with the metal center. The peak at ⬃378 nm has a relatively high extinction coefficient (ε 6000 M1 cm1) and was assigned to a S → Ni(III) LMCT [30]. A shoulder appears at 540 nm with a much lower extinction coefficient, and was initially assigned to a d-d transition. A detailed analysis of the electronic absorption spectrum using MCD reveals a more complicated spectrum made up of ten transitions (Figure 7) [82]. Variable-temperature variable-field MCD confi rm the S 1/2 electronic configuration. INDO/S-CI calculations were used to assign two bands responsible for the intense feature near 378 nm to S(σ) → Ni(III) d x2y2 transitions (Table 3). The amide N(σ)→Ni(III) LMCT also contributes to the observed optical

Table 3. Spectral parameters from NiSODox and NiSODred as well as transition assignments. NiSODox band 1 2 3 4 5 6 7 8 9 10 NiSODred band A B C D E F G

νmax (cm1)

εabs (M1 cm1)

Assignment

14 380 15 650 18 000 19 920 23 290 24 700 26 850 28 720 30 400 31 950

300 510 1340 1510 2020 3080 6800 4020 4900 4740

Ni(z2) → Ni(x2y2) S/N/O (π) → Ni (σ) S/N/O (π) → Ni (σ) S/N/O (π) → Ni (σ) S/N/O (π) → Ni (σ) N/O(σ) → Ni(x2y2) S6 (σ) → Ni(x2y2) S2 (σ) → Ni(x2y2)

νmax (cm1)

εabs (M1 cm1)

Assignment

17 110 18 430 20 500 22 240 24 970 27 650 29 220

70 150 180 480 500 880 750

Ni(xz/yz) → Ni(x2y2) Ni(xz/yz) → Ni(x2y2) Ni(z2) → Ni(x2y2) Ni(xy) → Ni(x2y2) S (σ / π) → Ni(x2y2) S (σ / π) → Ni(x2y2) S (σ / π) → Ni(x2y2)

Extinction coefficients for NiSODox are based on a 50:50 mix of oxidized:reduced enzyme and are corrected for by subtraction of the NiSODred spectrum. The transition assignments are the major contributions. Table adapted from [82].

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Figure 7. Absorption, CD, and MCD spectroscopy of NiSOD. Top panel, as-isolated NiSOD: low-temperature electronic absorption spectrum (top), CD spectrum (middle), 7 T MCD spectrum (bottom). Dotted lines represent the minimum set of Gaussian shaped transitions needed to simulate (dashed) the observed spectra (solid). Bottom panel, dithionite-reduced NiSOD: electronic absorption spectrum (top) and CD spectrum (bottom). Reproduced by permission of the American Chemical Society from [82]. Met. Ions Life Sci. 2, 417–444 (2007)

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absorption. The d z2 → d x2y2 transition was assigned to features at ⬃700 nm. The d xy → d x2y2 transition contributes to the middle range of the spectrum (near 450 nm) along with a number of S and N π → Ni(III) LMCT transitions. Upon reduction of the Ni(III) center, a nearly featureless optical absorption spectrum is obtained. However, MCD shows seven transitions between 700 and 300 nm. The bands are generally blue-shifted in the reduced enzyme, with the d z2 → d x2y2 and d xy → d x2y2 transitions shifting to ⬃488 nm and ⬃450 nm, respectively (Figure 7, Table 3). Resonance Raman spectra of the as-isolated enzyme exhibit three vibrations, two intense bands at 349 and 365 cm1 and a weaker band at 391 cm1. The excitation wavelengths used, 413 and 407 nm, are near that of the intense charge-transfer band associated with the S → Ni(III) electronic transition. This data was used to assign the intense bands at 349 and 365 cm1 to Ni(III)-S stretching modes involving the cysteine residues. The transition at 391 cm1 was assigned to either a bending frequency of the Cys2-S to Cys2-N chelate along with Ni-S stretching or a Ni-NCys2 stretching mode of the backbone deprotonated amide that is intensified by the charge-transfer transitions.

4.2.

Kinetics

The kinetics of the NiSOD reaction have been examined using pulse-radiolytic generation of superoxide (Figure 1) [30]. Pulse radiolysis offers a direct method of measuring the rates of the SOD reaction. Superoxide is generated quantitatively, and its absorbance at 260 nm is monitored on a rapid time scale, spanning milliseconds to seconds. Measured rate constants are based on metal concentrations to better compare the various classes of SOD. Studies of the NiSOD reaction kinetics establish that the enzyme operates by the ‘ping-pong’ mechanism that has been described for other SODs (Equations 1–3), at a rate near the diffusion limit (kcat ⬃ 109 M1 s1), and exhibits a pH dependence remarkably similar to MnSOD despite the lack of any sequence homology and the strikingly different active site (Figure 1). NiSOD does not have an aqua/hydroxo ligand, and so the mechanism exhibited by the Mn/FeSODs for coupling the protonation of hydroxide to the reduction of the enzyme, or for tuning the Ni(III/II) redox potential, is not available for the nickel enzyme. In spite of the absence of aqua/hydroxo ligands for NiSOD, residues that are analogous to Tyr34 in the Mn/FeSOD enzymes are present in the NiSOD active site (Tyr9, Tyr62) and participate in an H-bonding network involving water molecules and other residues that could serve as a source of protons. Another potential role for the tyrosine ligands as radical scavengers has also been suggested [62]. Like the CuZnSODs, NiSOD also has an imidazole ligand that is lost upon reduction [73,79], and a mechanism that is reminiscent of the CuZn enzyme, but does not Met. Ions Life Sci. 2, 417–444 (2007)


435 Cys2 N H

O

HO

O2–

NH

H2O2

S

N Ni (III) S NH2 N

Cys6 H+

O

Tyr 9

His1 Cys2 N H

O -

N H Cys6

S

N H

Cys2

HO

O

N Ni (II) S NH2 HN

O

O

O

N H

O Glu17

O

S

His1

O

Tyr 9

His1

N N H

H O

HO

O

N Ni (III) S NH2 N

Cys6

Tyr 9

O -

NH

O O

Cys2 Glu17

N H

H+ O

Glu17

HO

NH Cys6 O2–

S

N Ni (II) NH2 S

O

O2

O

Tyr 9

His1

HN N H O

O

Glu17

Figure 8. A hypothetical inner-sphere mechanism for NiSOD catalysis illustrating the putative role of the His1 imidazole ligand and Glu17 in redox tuning and/or H transfer.

involve imidazolate [82], is possible (Figure 8). This mechanism (described as an innersphere reaction although no experimental data regarding the innersphere versus outersphere character of the reaction has been reported) illustrates the H-bonding interaction with Glu17 that is a property of the hexameric enzyme. Mutations of the apical His1 imidazole ligand establish its important role in the catalytic mechanism. H1Q-NiSOD is isolated in a mostly reduced, Ni(II) state, suggesting that the redox chemistry is perturbed by this mutation. The enzyme still turns over by the ping-pong mechanism, but at a rate that is two orders of magnitude slower than the native enzyme [76]. Other candidates for proton donors Met. Ions Life Sci. 2, 417–444 (2007)

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include the thiolate ligands, which appear to pick up protons upon reduction by H2O2, but not upon reduction by other methods [61]. The ionic strength dependence of kcat has been measured for several SODs. In particular, catalysis in CuZnSOD shows a significant ionic strength dependence that has been attributed to the involvement of lysine in enhancing the electrostatic steering of superoxide to the active site of the enzyme for processing [83]. Relative to the effects seen in the CuZn enzymes, NiSOD kinetic data show very little ionic strength dependence [30], suggesting that very little electrostatic steering is involved in the mechanism.

4.3. Model Studies The spectroscopic studies coupled with the crystallographic data facilitated the development of theoretical models of the nickel active site that further advanced the understanding of the structure and reaction mechanism of NiSOD. Calculations on the Streptomyces sp. NiSOD active-site [82] were based on the crystallographically determined coordinates [73,79] employing the Amsterdam Density Functional software package. Constraints were employed based on assumptions that the protein determined a limited range of possible positions the backbone atoms and second sphere residues could possess. Also, modifications to the structure were made treating Cys6 as ethanethiolate, and removing the backbone carbonyl from Cys2. The B3LYP hybrid-density functional method in conjunction with the triple-ζ basis set (TZV) was used for the calculations of spin restricted systems in the case of Ni(II) and spin-unrestricted for Ni(III). Active-site models generated with DFT calculations reproduce structural parameters of the known NiSOD structures within 0.05 Å for the equatorial ligands. In particular the calculated states referred to as ox1 and red by Fiedler et al. [82] reproduce metric values very close to the crystal structures of the oxidized and reduced forms of NiSOD. However, difficulties were encountered in predicting the apical Ni–N bond distance, which was initially calculated to be 2.07 Å, well short of the crystallographically determined value of 2.35–2.63Å [73,79]. Hydrogen bonding interactions were examined between His1 and Glu17 and extended to include the interaction between Glu17 and Arg47. The incorporation of these additional interactions lengthened the axial Ni–N bond to at most 2.16 Å. While this improved the overall structural model of the NiSOD active site, it did not reproduce the much longer crystallographically determined distance. Deprotonation of the apical imidazole to give an imidazolate ligand was considered, but resulted in even shorter distances. Two possibilities factor into the 0.2 Å discrepancy: either the truncations and the constraints of the active site imposed on the calculations inadequately describe the active site, or the inhomogeneous oxidation state of the metal centers in the NiSOD crystals used for X-ray analysis generated an erroneously long Ni–N distance for the apical imidazole. Nonetheless, the calculations reveal that Met. Ions Life Sci. 2, 417–444 (2007)


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hydrogen bonding interactions with amino acid residues in the second coordination-sphere influence the axial bond distance of the His1 N-donor ligand to a large extent, and therefore play an important role in optimizing the redox properties of the active site. The model studies predict that the orbitals used for the two half-reactions in the ping-pong mechanism are different: Ni dz2 -based for the oxidation of O 2 , and involving a π-symmetry orbital for the reduction of O . The best model shows 2 that the unpaired electron in oxidized NiSOD resides in a MO that is largely a dz2 orbital that has 6% N character from the apical imidazole N-donor atom and is 76% Ni in character, consistent with a Ni(III) formulation and the interpretation of the EPR spectrum given above. The large N character accounts for the large hyperfine observed. The apical imidazole ligand plays an important role in optimizing the redox potential, as varying the strength of the apical Ni–N interaction by varying the Ni–N distance from 2.07 to 2.16 Å was shown to lower the energy of the Ni dz2 orbital by 0.23 eV without greatly affecting the energies of other orbitals. The pyramidal geometry of the Ni(III) site with strong basal σ-donors leads to a large destabilization of the Ni d x2y2 orbital. This is due in large part to the covalent bonding between Ni and the two cysteinate ligands, and results in a MO that has roughly 50% Ni and 30% S character. The basal S-donors and the amide N-donor both feature π-symmetry orbitals that align with the dz2 orbital to contribute significant π-conjugation. Upon reduction to Ni(II), the apical imidazole ligand is lost. The loss of the apical ligand stabilizes the Ni dz2 -based MO, and the decrease in positive charge on the Ni center destabilizes the π-symmetry d orbitals, resulting in Ni- and ligand-based orbitals with more similar energies, and thus higher covalency in the Ni(II) site. As a result, the HOMO is a π-symmetry orbital that is roughly 30% Ni and 60% S in character. Despite the ligand-based character of the HOMO in the planar Ni(II) complex, π-antibonding interactions from the S-donors and amide N-donor destabilize the π-symmetry orbitals in the Ni(III) complex, resulting in Ni-based oxidation (a Mulliken population analysis of the planar Ni(III) model showed that 70% of the unpaired spin density resides on the Ni center). It was proposed that the amide N-donor plays an important role in stabilizing Ni redox over S-centered redox. The possibility that protonated thiolates coordinated to the metal active site in NiSOD might serve as a source of protons for H2O2 production was also examined computationally. Individual models were geometry optimized that included either a protonated Cys2 or Cys6 thiolate. The resulting structures showed little effect by addition of the proton, and in fact produced shorter Ni–S bond lengths. This is consistent with model studies that involve protonation [84], alkylation [56,85– 87], and oxidation [36,37,53,55, 88–91] of sulfur in nickel thiolate complexes and reveal that little structural change accompanies the chemical modifications. The computational results correlate well with the known model complexes and support the model for coordinated thiols in reduced NiSOD. Met. Ions Life Sci. 2, 417–444 (2007)

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5.


CONCLUSIONS

The understanding of NiSOD structure and function that emerges from the data is one that suggests convergent evolution. The NiSOD protein has a primary structure that bears no resemblance to any other class of SOD. Yet, NiSOD appears to catalyze the same reaction as other SODs, with the same remarkable efficiency, and using a ping-pong mechanism involving a metal that oscillates between oxidized and reduced states. It does so using a unique ligand environment that creates a novel nickel center where the Ni(III/II) redox couple is lowered by over 1V. A combination of several strategies appears to be employed in NiSOD to achieve the requisite redox potential. The enzyme features a ‘nickel hook’ domain in the N-terminus of the protein that requires post-translational processing and becomes structurally robust only upon Ni incorporation. Three residues in that domain contribute all of the nickel ligands, which includes two thiolate ligands and an amidate N-donor that lower the redox potential of the system and favor metal-centered over ligand-centered redox processes. Mutagenesis has established a critical role for the apical imidazole ligand, whose role in tuning the nickel redox potential is clearly defined by theory. The source of protons that are required for the production of H 2O2 is not clear, though it is apparent that the mechanisms used by the Mn/Fe and CuZn enzymes do not adequately describe the situation for the Ni site. Although a H-bonding network involving residues in the second coordination sphere (such as tyrosine residues and water molecules) like that found in the Mn/Fe enzymes is present, the lack of an aqua/hydroxo ligand rules out a mechanism that is exactly analogous to those enzymes. Similarly, in parallel with CuZnSODs, the NiSOD active site loses an imidazole ligand upon reduction. However, this ligand is an imidazole, not an imidazolate, and thus cannot serve as a protonation site in the same way as the bridging imidazole in the CuZn site without the involvement of residues in the second coordination sphere. In addition, S K-edge XAS and theory have provided evidence supporting a role for the cysteinate ligands in supplying protons for catalysis. Although NiSOD takes advantage of many features found in other SODs, the use of a metal that lacks a natural redox chemistry in water, and the application of the protein to create an appropriate active site, represent a completely novel approach to the elimination of superoxide in biological systems, and thus contribute to the biodiversity of mechanisms for oxygen detoxification.

ACKNOWLEDGMENTS We thank Dr Diane Cabelli for helpful comments on this manuscript and acknowledge support from the Samuel F. Conti Faculty Fellowship (University Met. Ions Life Sci. 2, 417–444 (2007)


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of Massachusetts, M.J.M.) and the National Science Foundation (MCB-021482, M.J.M.). Trainee funding (P.A.B.) was provided by the NIH-Chemistry-Biology Interface Program (T32 GM08515).

ABBREVIATIONS AND DEFINITIONS CD CODH CT CuZnSOD cyclam DTT DFT EPR ESI-MS EXAFS fALS FeSOD HiPIP HOMO IMAC INDO/S-CI LMCT MCD MnSOD MO NHE NiSOD PCET ROS rR SOD XANES XAS

circular dichroism carbon monoxide dehydrogenase charge-transfer copper zinc superoxide dismutase 1,4,8,11-tetraazacyclotetradecane dithiothreitol density functional theory electron paramagnetic resonance electrospray ionization mass spectrometry extended X-ray absorption fine structure familial amyotrophic lateral sclerosis iron superoxide dismutase high-potential iron protein highest occupied molecular orbital immobilized metal affinity chromatograph intermediate neglect of differential overlap/single-configuration interaction ligand to metal charge-transfer magnetic circular dichroism manganese superoxide dismutase molecular orbital normal hydrogen electrode nickel superoxide dismutase proton-coupled electron transfer reactive oxygen species resonance Raman superoxide dismutase X-ray absorption near-edge structure X-ray absorption spectroscopy

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11 Biochemistry of the Nickel-Dependent Glyoxalase I Enzymes Nicole Sukdeo, Elisabeth Daub, and John F. Honek* Department of Chemistry, University of Waterloo, 200 University Avenue, Waterloo, Ontario, N2L 3G1, Canada [email protected] [email protected] [email protected]

1.

INTRODUCTION 1.1. Methylglyoxal Formation 1.2. Cellular Effects of Methylglyoxal 1.3. Methylglyoxal Degradation and the Glyoxalase System 2. BIOCHEMICAL INVESTIGATIONS OF GLYOXALASE I 2.1. Survey of Glyoxalase I Enzymes from Various Sources 2.2. Metal Activation of Glyoxalase I Enzymes 2.3. Two Classes of Glyoxalase I 3. BIOPHYSICAL AND MECHANISTIC STUDIES OF GLYOXALASE I 3.1. Biochemical Investigations of Glyoxalase I 3.2. Biophysical Investigations of E. coli Glyoxalase I 3.3. Glyoxalase I-Catalyzed Isomerization and Proposed Mechanism

*

446 446 448 449 450 450 451 452 453 453 455 458

The authors wish to dedicate this chapter to Professor Donald Creighton, who passed away in early 2006. His many contributions to the glyoxalase field as well as his kind friendship are greatly appreciated.



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4.

GLYOXALASE I GENES AND PROTEIN SEQUENCE COMPARISONS 4.1. Relationship of Sequence to Metal Activation Profile 5. GLYOXALASE I AS A MEMBER OF THE βαβββ SUPERFAMILY OF PROTEINS 6. OTHER ASPECTS OF GLYOXALASE I 7. CONCLUSIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

1.

460 460 463 464 465 466 466 467

INTRODUCTION

In 1913, two groups independently reported their discovery of enzymatic production of D-lactate and D-mandelate from methylglyoxal (MG) and phenylglyoxal, respectively, in eukaryotes [1–3]. However, it was 37 years before it was realized that two enzyme activities, glyoxalase I (GlxI) and glyoxalase II (GlxII) (Figure 1) were responsible for formation of these products [4,5]. Nevertheless only recently have the molecular structures and differing metal specificities of these enzymes been determined. This review will focus on the GlxI enzymes and the identification of two distinct classes of this enzyme, a zinc-dependent as well as a non-zinc-dependent (maximal activation with nickel) class which appear critical for detoxification of cellularly produced MG.

1.1.

Methylglyoxal Formation

A direct enzymatic route for production of methylglyoxal has been known to exist in members of the Enterobacteriaceae [6]. MG synthase was first identified in Escherichia coli and catalyzes MG formation from dihydroxyacetone phosphate (DHAP). This enzymatic activity was first inferred through the observation of MG accumulation in triose phosphate isomerase-deficient mutants of E. coli [7]. Cooper and Anderson proposed that conversion of MG to pyruvate, via the enzymes MG synthase, the glyoxalase system and lactate dehydrogenase, acting in tandem, provides a metabolic scheme for pyruvate production by bypassing its glycolytic route of formation (Figure 2) [7]. Accumulation of inorganic phosphate (Pi) was observed to inhibit MG synthase, lending support to the idea that the glycolytic bypass was activated Met. Ions Life Sci. 2, 445–472 (2007)

NICKEL-DEPENDENT GLYOXALASE I ENZYMES

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O O

+H N 3

O

H

Non-Enzymatic

NH

CH3 +

N H

CO2–

O

CH3

H

SH γ-glutamyl-L-cysteinylglycine (GSH)

Methylglyoxal (MG)

O

GS HO

CO2–

Hemithioacetal GlxI

O

O– CH3

GS HO H -D-Lactoylglutathione GlxII

O

GS

OH OH GS CH3

O– CH3

H2O

H

OH CH3 D-Lactate

–O

Figure 1. Reactions of the glyoxalase system. The proposed enediol(ate) intermediate for the GlxI-catalyzed reaction is shown above.

Fructose-1,6-Bisphosphate (FBP)

GSH DHAP

Glyceraldehyde-3Phosphate

MG

Hemithioacetal

Phosphorylating Glycolysis

Pyruvate

GSH + D-Lactate

Figure 2. The glyoxalase system as a bypass of phosphorylating glycolysis. TIM: triose phosphate isomerase. Met. Ions Life Sci. 2, 445–472 (2007)

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under conditions of phosphate limitation in the cell. Therefore, if the cytosolic pools of DHAP are high relative to pools of Pi, MG synthase activity would increase, permitting pyruvate formation through the non-phosphorylating enzymatic bypass. This re-routing of DHAP consumption would preserve the existing pool of Pi, permitting phosphorylating glycolysis when the Pi concentration was sufficient to induce glyceraldehyde-3-phosphate dehydrogenase activity [8]. The glycolytic enzyme triose phosphate isomerase (TIM) is another documented and potentially substantial means of MG production in vivo [9]. Normally responsible for catalyzing interconversion between dihydroxyacetone phosphate and glyceraldehyde-3-phosphate, the enediolate intermediate generated during catalysis by TIM is susceptible to inorganic phosphate elimination. This alternative reaction results in MG formation and the frequency of elimination occurring is estimated to occur once in approximately 100 000 turnovers by the enzyme [10]. This frequency of TIM proceeding through elimination can translate into a rate of MG formation in cells approaching 0.4 mM/day [9]. This substantial rate of accumulation of MG is due in part to the typically high cellular concentrations of triose phosphate isomerase in metabolically active cells, underlining the need for constitutive routes of detoxification [9,11]. Amino acid degradation provides yet another route for MG production. More specifically, glycine and threonine degradation yields aminoacetone from which MG is obtained following conversion by monoamine oxidase [12–14]. The existence of amine oxidases that produce MG has been confirmed in Saccharomyces cerevisiae and Staphylococcus aureus [12]. Isozymes of the cytochrome P450 IIEI subfamily are also capable of producing MG from acetone [15,16]. The discovery of acetone-converting monooxygenases resulted from analysis of hepatic microsomes from rats that consumed acetone in drinking water. The analysis revealed the presence of an acetone monooxygenase that converts acetone to acetol. A second acetol monooxygenase converted the acetol to MG [15].

1.2. Cellular Effects of Methylglyoxal Staehelin discovered that 2-oxoaldehydes are capable of interacting with and inactivating tobacco mosaic virus RNA, particularly by interaction with guanidine derivatives [17]. The electrophilic nature of α-ketoaldehydes was established through investigations of the toxicological properties of MG; MG and other α -ketoaldehydes are capable of forming adducts with nucleic acids and proteins. MG accumulation is considered to exert deleterious effects at several levels of cellular function, even though the precise mechanism of toxicity still remains unclear [8,18–20]. Lipid modification and cell death from oxidative damage, in addition to increased apoptosis are among the diverse cytotoxic effects that have been associated with MG in the recent literature [21–24]. Met. Ions Life Sci. 2, 445–472 (2007)


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MG can inhibit the synthesis of protein and RNA by forming adducts with these classes of macromolecules. Fraval and McBrien observed almost immediate inhibition of protein synthesis in E. coli with the addition of MG, compared with a delay in inhibition of DNA synthesis [25]. It was concluded that the primary cytotoxic effect of MG was the inhibition of protein synthesis. The selective inhibition of DNA synthesis at initiation was attributed to MG interaction with the initiation complex or possibly inhibition of the synthesis of proteins involved in the initiation of DNA replication [25]. MG adducts of nucleic acids are genotoxic, either directly or through potentiation of oxidative damage [26–28]. Carbonyl compounds, including MG, can cause a number of metabolic changes in the cell. These compounds can engage in non-enzymatic, irreversible glycation of amino acids [29]. The advanced glycation end products (AGEs) found in nucleic acid and protein molecules result from a series of reactions, involving rearrangements, dehydration, and redox chemistry, making the precise mechanism of covalent modification rather complex [29]. In addition, little is known concerning the direct effects of the product of GlxI, S-lactoylglutathione. Elevated levels of MG have been associated with states of hyperglycemia, and overexpression of GlxI has been observed to minimize the amount of AGEs formed in cells based on in vitro studies [30,31]. In addition to the α-ketoaldehydes, physiological effects of the GlxI product (S-lactoylglutathione) have been investigated. The ability of S-lactoylglutathione to elicit histamine from mast cells has been reported in the literature [32–34]. However, the main focus of α-ketoaldehyde biological effects has been the toxicity of MG and related derivatives. This research also examined the effects of the inhibition of GlxI which is expected to result in an increase in the levels of cytotoxic MG in cells. Subsequently small molecule inhibitors of human GlxI have been developed and have shown interesting cellular toxicity in select cancer cell lines, due to their possibly higher glycolytic activity and resulting higher requirements for MG metabolism to avoid toxicity [35–38].

1.3.

Methylglyoxal Degradation and the Glyoxalase System

The need for detoxification of MG and other α-ketoaldehydes can be accomplished in a number of different ways. NADPH-dependent oxidoreductases that catalyze conversion of MG to acetol or lactaldehyde have been identified in E. coli, S. cerevisiae, and several mammalian species. MG conversion to lactaldehyde by MG reductases has been characterized in S. cerevisiae and Aspergillus niger and has also been identified in mammalian liver homogenates [39–41]. Additionally aldose reductase enzymes have been characterized as another route of MG catabolism [30,42]. Homologs of mammalian aldo–keto reductases have been identified in E. coli that accept MG and phenylglyoxal as substrates for NADPH-dependent reduction [43]. Gram-positive organisms lacking GSH can rely on other metabolic enzymes for MG detoxification. In the case of some Met. Ions Life Sci. 2, 445–472 (2007)

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Clostridia species, glycerol dehydrogenase activity is principally associated with removal of cellular MG [44]. This dehydrogenase is thought to decrease cellular MG concentrations by reduction of dihydroxyacetone [44]. Another enzyme system which appears to make a major contribution to metabolism of MG is the glyoxalase system, and in fact, is believed to be the major enzyme system responsible for MG detoxification [6]. Certainly, the glyoxalase system can serve as a direct detoxification system such that MG can be converted to innocuous D-lactate. Interestingly, it has also been shown that for E. coli, if MG concentrations reach certain levels (0.3–0.6 mM), a buildup of S-lactoylglutathione may occur with concomitant reduction in GSH levels due to the differential rates of GlxI versus GlxII activity. An important set of efflux pumps, KefB and KefC are affected negatively by GSH levels and positively by S-lactoylglutathione levels. Activation of the KefB and KefC potassium efflux systems present in E. coli results in concurrent influx of Na and H [45]. The reduction of intracellular pH from H influx correlated with reduction in MG toxicity, although the mechanism of this phenomenon remains as yet unknown. At low cellular concentrations of MG, available GSH inhibits K permeation through KefB and KefC. As the glyoxalase system converts MG to S-lactoylglutathione (by way of GlxI), increased levels of S-lactoylglutathione might arise which serve to activate KefB and KefC and thereby suggests a potential physiological role for this thioester in gram-negative bacteria [46]. Hence the important roles that the glyoxalase system plays in the control of MG levels, either directly or as a component of a whole-cell response to toxicity (Kef channels), have been the subject of much interest. Characterization of this metalloenzyme system has contributed further to the understanding of the role that metalloenzymes play in detoxification pathways. This chapter will focus on the first of the two enzymes, GlxI, concentrating on the structure and function of this enzyme.

2. 2.1.

BIOCHEMICAL INVESTIGATIONS OF GLYOXALASE I Survey of Glyoxalase I Enzymes from Various Sources

The glyoxalase system has been documented in a vast number of organisms as a metabolic pathway permitting conversion of methyglyoxal and other -ketoaldehydes to D-lactate or other -hydroxyacids, respectively. Occurrence of the glyoxalase system in nature spans eukaryal and prokaryotic organisms [47]. Two enzymes carry out the constituent reactions associated with this pathway (Figure 1). MG enters the glyoxalase system as a non-enzymatically formed hemithioacetal with GSH. GlxI (S-D-lactoylglutathione MG lyase (isomerizing) E.C. 4.4.1.5) is a metalloenzyme that converts the incoming hemithioacetal to the corresponding thioester, which from MG-GSH would be S-D-lactoylglutathione. This substrate Met. Ions Life Sci. 2, 445–472 (2007)


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is hydrolyzed by GlxII (S-2-hydroxyacylglutathione hydrolase E.C. 3.1.2.6), a binuclear metalloenzyme to generate D-lactate and GSH [48–53]. The substrate specificity of various GlxI enzymes has been investigated and it has been found that the human and rat enzymes are capable of handling large α-ketoaldehydes such as phenylglyoxal (and other arene-substituted glyoxals) and other larger aliphatic (and branched) ketoaldehydes in addition to MG [54]. The molecule 4,5-dioxovalerate has also been observed as a substrate for bovine liver and A. niger GlxI enzymes [55,56]. A variety of reports have appeared on the isolation and study of GlxI enzymes, to various levels of detail [47]. Putative GlxI sequences have also been identified and reported, underlining the broad occurrence of the enzyme system in Nature [57–68]. A search (as of January 2006) of the National Center for Biotechnology Information (NCBI) Protein Database reveals a diverse distribution of unique putative GlxI sequences: 70 bacterial sequences, 27 from the metazoan class, 34 fungal sequences, 8 from plant sources and 10 other eukaryotic sequences mainly from Plasmodium spp [69]. For example, the GlxI enzymes from Brassica juncea and soybean, and mouse have been reported in the literature [66,70,71]. However it has been the detailed studies of the GlxI enzymes from human, Saccharomyces cerevisiae, rat, and Pseudomonas putida that have yielded the most detailed information at the mechanistic level [72–77]. Recently, the cloning and expression of a number of other bacterial GlxI (E. coli, Pseudomonas aeruginosa, Neisseria meningitidis and Yersinia pestis) have contributed to additional and fascinating information about the GlxI enzymes [67,78,79].

2.2.

Metal Activation of Glyoxalase I Enzymes

A metal requirement for activation of GlxI was first demonstrated with the calf liver GlxI in 1966 by Davis and Williams [80]. Complete inactivation of the enzyme was noted after passage through a gel filtration column equilibrated with 2 mM ethylenediaminetetraacetate (EDTA). The authors observed a dosedependent increase in hemithioacetal turnover with MG with increasing concentrations of Mg2 using enzyme purified in the presence of EDTA. Other divalent metals were screened as cofactors for GlxI and several were found to be efficacious to varying degrees for reactivation of the enzyme. Several other mammalian GlxI enzymes were isolated and characterized in the 1970s, all displaying robust activation in the presence of Mg2 ions, and this was thought to be the activating metal in vivo for GlxI for a number of years [81–84]. Metal analysis of homogeneous GlxI from S. cerevisiae, porcine erythrocyte, human erythrocyte, and rat liver showed, however, that the naturally incorporated metal was Zn2 bound in a stoichiometry of one mole per subunit for the mammalian enzymes [85]. Human and rat liver GlxI apoenzymes also displayed significant reconstitution of activity in the presence of Zn2 ions and Met. Ions Life Sci. 2, 445–472 (2007)

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hyperactivation in the presence of Mg2 ions [85]. These findings indicated that GlxI occurs in nature as a zinc metalloenzyme, which was prevalent in many other GlxI enzymes characterized subsequently until study of the E. coli enzyme (see Section 3.1). Detailed metal reconstitution of the human enzyme also indicated a comparatively smaller dissociation constant for Zn2 relative to other divalent ions, in keeping with Zn2 activation as the natural metal [86]. No indication of non-zinc activation was reported; however, several studies have shown that the zinc-activated enzymes (P. putida and human) can in fact be demetallated to the inactive apoenzyme form and be remetallated with zinc ions or other metals such as cobalt, nickel, and magnesium, all of which confer various levels of enzymatic activity [77,85]. Another interesting feature realized from detailed study of the Zn2-activated GlxI enzymes is the ability of GlxI to isomerize both diastereomers of the α-ketoaldehyde-glutathione hemithioacetals [87–89]. The precise nature of how GlxI deals with these opposing configurations before or during catalysis still requires further investigation.

2.3. Two Classes of Glyoxalase I There are a number of reasons why zinc is a common metal found in a variety of metalloenzymes. For example, zinc is capable of alteration of its metal coordination and geometry during catalysis. Tetracoordinate, pentacoordinate, and hexacoordinate (as in GlxI) metal centers are prevalent in zinc metalloenzymes, and fluctuation of coordination geometry in the course of the catalytic cycle is a common mechanistic feature for these proteins [90]. Zinc centers also possess the capacity for rapid exchange of water ligands that may be analogous to on/off rates of incoming substrate (⯝107 s1) [91]. Zinc is most stable in its divalent form and is most stable in aqueous media, in addition to being prevalent in biological metal centers [92,93]. It also does not readily undergo reduction–oxidation reactions compared with other types of metal ions, (Mn, Co, Mo, etc) and its non-redox role is exemplified when considering GlxI enzyme catalysis. Zinc as a coordination center can exist stably in metal hydrate and metal hydroxide forms at physiological pH, permitting aqua ligands to be present in enzymes that have a capacity for general acid/base catalysis [93]. In addition, water molecules ligated to metal centers have altered pKa values which can render them hypernucleophilic and mechanistically relevant [90]. In addition to Ca2, Mg2, and Fe2, cellular Zn2 is accumulated to millimolar concentrations in cells (based on growth in complex and minimal media), but tight control is exerted over the free metal ion content in the cellular cytoplasm [94]. Such cellular concentrations highlight the remarkable capacity for metal ion sequestration relative to environmental abundance (10 −7 and 10 −5M Zn2 in minimal media and Luria Bertani (LB) media, respectively) [94]. Met. Ions Life Sci. 2, 445–472 (2007)


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The field of nickel metallobiochemistry to contrast, has a more recent history considering the timeline for identification of nickel enzymes relative to zinc activated enzymes. Ni2-metalloenzymes have been characterized less frequently and have been identified only within the last 30 years (urease identified in 1926, but nickel utilization discovered in 1975) [95–98]. Ni2 in biological function is associated with redox-active (methyl-CoM-reductase) as well as non-redox (urease) metallocentres [95,99–101]. Compared with Zn2-enzymes, anaerobic bacterial and archaeal species commonly contain Ni enzymes in their proteome [95]. As Ni-utilizing metalloenzymes continue to be discovered, as was the case with E. coli GlxI, a greater understanding of why this metal is desired in some enzymatic processes will be resolved to the extent achieved in the area of zinc metallobiochemistry. These studies on various zinc-activated GlxI enzymes suggested that newly isolated GlxI enzymes from new sources might also be zinc-activated. However, the surprising first report in 1998 of a non-zinc-dependent GlxI enzyme from the gram-negative organism E. coli indicated that perhaps this generalization is not correct [78]. This metal activation profile has recently been discovered in GlxI enzymes from P. aeruginosa, Y. pestis, and N. meningitidis, all displaying a propensity towards Ni2 activation, but no recovery of activity in the holo-Zn2 form [79]. The E. coli GlxI enzyme is representative of this novel class of metal activation owing to its extensive characterization. The biochemical features of this metalloenzyme are outlined in the following section.

3.

BIOPHYSICAL AND MECHANISTIC STUDIES OF GLYOXALASE I

3.1. Biochemical Investigations of Glyoxalase I The isolation of the E. coli enzyme provided a recombinant source for additional studies of a GlxI enzyme [78]. The E. coli enzyme was found to be dimeric, like the human enzyme, with a homodimeric quaternary structure as shown by sodium dodecylsulfate polyacrylamide gel electrophoresis (SDS-PAGE) in combination with gel permeation chromatography. The development of a purification scheme to produce substantial quantities of the enzyme (140 mg/L of cell growth) as well as a mild method to prepare demetallated apoenzyme was critical for further investigation of this system. With the availability of sufficient quantities of E. coli GlxI enzyme, it was possible to study in detail the interactions of various metals with this enzyme. Metal reactivation studies with the apoenzyme generated unexpected results, specifically, that Zn2 was incapable of reactivating this form of the apo form of the E. coli enzyme, but that cobalt and especially nickel were competent activating metals. Additionally, both Mn2 and Cd2 were also found to somewhat activate Met. Ions Life Sci. 2, 445–472 (2007)

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the E. coli GlxI. The Km ( µM) and Vmax ( µmol/min/mg) values for Ni2-, Co2-, Cd2- and Mn2-substituted enzyme have been determined and the following values have been obtained: (Ni2: Km 27, Vmax 676; Co2: Km 12, Vmax 213; Cd2: Km 8.9, Vmax 43; Mn2: Km 10, Vmax 121) [78,102]. In order to obtain information on whether Zn2 was capable of binding to the apo form at all, isothermal titration calorimetry (ITC) studies were pursued. Equilibrium Ka values for Ni2 and Co2 (both 107) association with apoenzyme correspond to dissociation constants in the nanomolar range (Kd ⯝108) for E. coli GlxI, whereas the Ka for Zn2 (Ka 108) is an order of magnitude below this value [102]. Ka values were also quantitatively obtained for titration of E. coli GlxI with Mn2 and Cd2 and yielded values of 3.9 ± 1.0 106 and 108, respectively [102]. Our preliminary differential scanning calorimetric (DSC) studies revealed that Tm values of Zn2–GlxI exceed those for the Ni2 and Co2 substituted forms by several degrees Celsius [102]. The apparent tightness of Zn2-binding and increased thermostability is intriguing, considering that it is not an activating metal for E. coli GlxI. However, these biophysical features in conjunction with pentacoordinate active site geometry (see below) may indicate a more rigid active site in Zn2-substituted E. coli GlxI, rendering this holoform catalytically incompetent. It is interesting to note that during the above-mentioned ITC studies, incubation of the E. coli GlxI apoenzyme in the ITC cell resulted in a slow, but measurable exothermic reaction in the absence of added metal from the titration syringe. For example, one hour of preincubation of the apoenzyme in the ITC cell without titration resulted in a reactivation of the enzyme to approximately 10% of its fully metallated form [102]. Metal analysis of this ‘apo’ form indicated that, in fact, nickel ion was being supplied from some source during the preincubation phase. Subsequent investigation determined that the cell, which is made of Hastelloy stainless steel which is 60% nickel in composition (Alfa Aesar 1999–2000 Research Chemicals, Materials and Metals), was apparently supplying the apoenzyme with Ni2 [102]. This resulted in detection of measurable exothermic heat as well as partial reactivation to a partially metallated form during this preincubation phase [102]. This experimental finding highlights the strong, and facile metal recruitment capability of the E. coli GlxI enzyme, which contrasts with reconstitution studies of the Zn2-dependent forms. The ease of metal removal and reintroduction into the E. coli GlxI dimer may be due in part to a more open structure compared with the human enzyme which demonstrates time-dependence of metal binding to the apo form. Reconstitution studies of human GlxI have shown that only partial reactivation is observed when reconstituting the apoenzyme, with Mg2 being a better activating metal under such conditions compared with Zn2 [85]. Mg2 is not an activating metal for E. coli GlxI, and our ITC studies give no indication that this ion binds to the enzyme active site [102]. The S. cerevisiae GlxI enzyme is completely recalcitrant to metal activation following treatment with chelating agents, in keeping with the reconstitution limitations observed for human GlxI [85]. Met. Ions Life Sci. 2, 445–472 (2007)


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3.2. Biophysical Investigations of E. coli Glyoxalase I Several biophysical studies on the metal center of the Zn2-activated human GlxI have been reported [103–107]. These investigations were valuable for developing a hypothetical model of the GlxI active site well in advance of the first crystal structure for this enzyme (human GlxI) in 1997 [108]. Human GlxI enzyme has been characterized with a paramagnetic metal centre (Mn2, Co2), allowing for investigation through electron paramagnetic resonance, and nuclear relaxation studies [104,109]. These studies indicated that the metal center possesses an octahedral geometry with no direct coordination of GSH, S-D-lactoylglutathione or S-p-bromobenzylglutathione to the active site metal [104,109]. Biophysical characterization of the E. coli GlxI active site (X-ray absorption spectroscopy, X-ray crystallography) has also generated a similar picture for the Ni2-activated enzyme, also possessing octahedral geometry and indirectly associating with substrate/product molecules/analogs [110–112]. A detailed structural characterization of E. coli GlxI has been accomplished over the past few years, permitting comparison with Zn2-activated GlxI, and the details of these studies are summarized below. Extended X-ray absorption fine structure (EXAFS) and X-ray absorption nearedge spectroscopy (XANES) studies of the Ni2 and Zn2 bound forms of the E. coli enzyme indicated that the active site geometry surrounding the nickel ion was most likely octahedral (six-coordinate) and that the zinc-containing form of the enzyme was different, likely five-coordinate in a trigonal bipyramidal arrangement [110,112]. Ni2-coordination was not modified in the presence of S-lactoylglutathione which appears similar to what was observed for nuclear relaxation studies of the human GlxI as mentioned previously [104,112]. Imidazoleand carboxylate-containing side chains were predicted through interpretation of EXAFS data to comprise the metal active site of E. coli GlxI [110]. Crystallographic studies on the E. coli GlxI enzyme were also undertaken and proved invaluable in elucidating the structural properties of the E. coli enzyme. Similar to the human GlxI enzyme, E. coli GlxI was dimeric in nature with two active sites that were situated at the subunit interface [108,111]. The structure also revealed that E. coli GlxI possesses a similar overall fold to that of the human GlxI, such that both enzymes are members of the βαβββ superfamily (see Section 5) [111,113]. The overall structures of GlxI enzymes from E. coli and human overlap well, with almost identical positioning of the active site metal ligands in spite of the difference in amino acid sequence length between these proteins (Figure 3 and Figure 6 below) [108,111,113]. The human enzyme possesses an extended N-terminal arm (29 amino acids in length), which is a common feature of other Zn2-dependent GlxI enzymes (Figure 3 and Figure 6 below). The active site of human GlxI is smaller than that of E. coli GlxI, constrained by a 15-amino acid loop helix that forms a posterior wall to a 70 Å3 hydrophobic pocket in the former enzyme [111]. In contrast, the E. coli GlxI active site is a deep solvent channel, Met. Ions Life Sci. 2, 445–472 (2007)

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Figure 3. Structures of: (A) Ni2-bound E. coli; (B) Zn2-bound human GlxI enzymes. Each monomeric subunit in the GlxI dimer is indicated by differential shading. Enlargements of the active site for both enzymes are shown to the right of the dimers with the bound metal represented as a dark sphere.

10–15 Å in diameter. This size discrepancy may reduce restrictions on the size of possible substrates for the E. coli enzyme. E. coli GlxI has been successfully crystallized as an apoenzyme in addition to Zn2-, Ni2-, Co2-, and Cd2-reconstituted forms [111]. No major structural changes accompanied metallation of the apoenzyme, corroborating the absence of secondary structure alteration between apo versus holo states as observed for E. coli GlxI in circular dichroism experiments [78]. The dimeric assembly of the enzyme was unchanged comparing apoenzyme with metal-reconstituted E. coli GlxI, based on the elution profiles of these various forms during gel filtration chromatography [78]. Crystallographic studies have shown that the various metal-activated forms of E. coli GlxI all possess octahedral geometry around the activating metal with four of the metal ligands being supplied from both subunits of the protein (His5, Met. Ions Life Sci. 2, 445–472 (2007)


457

Figure 4. Coordination of (A) Ni2 and (B) Zn2 in E. coli GlxI. The water O/OH spheres represent ligation of oxygen atoms to the metal center.

Glu56 from one monomer and His74, Glu122 from the other) as well as two water (or hydroxide) ligands completing this geometry (Figure 4) [59, 111]. Attempts to produce crystals of the Mn2-containing enzyme were not successful, although electron spin resonance experiments were consistent with an octahedral environment for this activating metal [111]. Interestingly, the zinc metallated form of the enzyme was found to have a very different metal geometry, a five-coordinate (trigonal bipyramidal) metal center possessing all amino acid ligands and only one water/hydroxide ligand (Figure 4) [111]. It appears Met. Ions Life Sci. 2, 445–472 (2007)

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from these studies that the activated form of the enzyme requires an octahedral geometry around the metal site with two water molecules as additional ligands to those supplied by the protein. Perhaps this arrangement is important in the polarization of substrate and in some way, the deprotonation of the substrate to form the enediol(ate) intermediate with subsequent reprotonation to form the product, S-lactoylglutathione (Figure 1) [110,112]. In any case, the octahedral metal environment correlates to active metal centers from both the human and E. coli GlxI, suggesting that this geometry might be a prerequisite for catalysis in Zn2 and non-Zn2-dependent enzymes. The stoichiometry of metal-binding in E. coli GlxI presents an interesting conflict between X-ray structure data and other biochemical studies of the enzyme. Titration of the apo-GlxI with increasing equivalents of Ni2 and Co2 restores maximal activity (for the given metal ion) when the stoichiometry is close to one metal ion per dimer. Although the enzyme is a homodimer with two active sites, metal analysis by inductively coupled plasma atomic emission spectroscopy (ICP-AES) and the ITC study of enzyme reconstitution also indicate that the Ni2 and Co2 reconstituted GlxI possess one equivalent of metal per dimer [78,102]. However, X-ray structures of the Cd2-, Ni2-, Co2-, and Zn2-reconstituted enzyme exhibit full occupancy of both active sites [111]. It is possible that complete metallation of the homodimer was the only form of the holoenzyme amenable to proper crystal formation. We are actively pursuing additional studies to firmly establish whether there is a differential affinity of each active site for metal through nuclear magnetic resonance (NMR) spectroscopy. This approach should allow us to probe microscopic structural changes accompanying titration of metal into the apoenzyme, establishing whether the data from metal analysis and ITC correlates to microscopic asymmetry of the dimer interfaces.

3.3. Glyoxalase I-Catalyzed Isomerization and Proposed Mechanism Based on a substantial amount of mechanistic analysis of the yeast and mammalian enzymes the likely mechanism for GlxI appears to involve an enediolate intermediate generated by enzyme catalyzed proton abstraction (Figure 1) [74,89,114– 116]. The structures of the human enzyme in the presence of S-benzylglutathione, S-p-nitrobenzyloxycarbonylglutathione (NBC-GSH) (Figure 5) and the hydroxamate S-(N-hydroxy-N-p-iodophenylcarbamoyl)glutathione (HIPC-GSH) (Figure 5) have been obtained providing some insight into the nature of substrate binding to human GlxI [108,117]. NBC-GSH structurally resembles the GlxI thioester product, but S-(N-aryl-N-hydroxycarbamoyl)glutathione derivatives are tightbinding inhibitors which were designed as mimics of the enediolate intermediate postulated to form during the enzymatic reaction [118,119]. Met. Ions Life Sci. 2, 445–472 (2007)


459 NO2

(A) O S

O +H N 3

H N

N H

CO2–

O

CO2–

O OH

(B)

O S

O +H N 3

H N

N H

CO2–

N I CO2–

O OH

(C) H3C

N

O

O +H N 3

CO2–

N H

H N

CO2–

O

Figure 5. Chemical structures of: (A) S-p-nitrobenzyloxycarbonylglutathione (NBCGSH); (B) S-(N-hydroxy-N-p-iodophenylcarbamoyl)glutathione (HIPC-GSH), and (C) L-glutamyl-N-hydroxy-N-methyl-L-glutaminylglycine.

The complexes of human GlxI with S-benzylglutathione and NBC-GSH were useful in probing conformational changes in the glutathionyl moiety of the substrate upon binding. However the HIPC-GSH-bound GlxI exhibited changes in metal coordination that were pertinent in formulating a hypothetical catalytic mechanism for the enzyme. The human GlxI-HIPC-GSH co-crystal structure exhibited displacement of one of the metal ligands (Glu173) [117]. It is therefore possible that this displaced Glu could serve in the role of a general base for proton abstraction at some point along the reaction pathway. This observation was intriguing given the lack of a clearly defined general base in the active sites of either the human or the E. coli enzyme upon inspection of both crystal structures [108,111]. This possibility has even been suggested by computational modeling of the zinc-containing human active site [120,121]. Studies of a similar hydroxamate, L-glutamyl-N-hydroxy-N-methyl-L-glutaminylglycine (Figure 5) in the case of E. coli GlxI utilizing EXAFS methods have suggested a similar displacement Met. Ions Life Sci. 2, 445–472 (2007)

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of a carboxylate ligand (Glu122) from the nickel center [122]. The similarities in structure for the E. coli and human GlxI-hydroxamate complexes are therefore suggestive of a mechanistic attribute shared by both metal activation classes. Nevertheless, caution is required in interpreting the aforementioned studies. Although hydroxamates have been utilized to mimic the reactive intermediate enediolate for the enzyme triose phosphate isomerase, they have also served as useful probes in determining isomerization mechanisms [10]. These inhibitors have been employed in the elucidation of the accepted hydride transfer mechanism xylose isomerase [10,123]. As well, and perhaps most importantly, hydroxamates have a long history of use as chemical moieties important in metalloprotease inhibition [124]. Since metalloproteases do not utilize an enediol(ate) intermediate in their mechanism, it is clear that metal-binding affinity of these types of compounds can override their subtlety in mechanistic studies.

4. GLYOXALASE I GENES AND PROTEIN SEQUENCE COMPARISONS 4.1.

Relationship of Sequence to Metal Activation Profile

A key aspect to our understanding of GlxI enzymes has been the discovery of two separate metal-activation classes. Establishment of these classes has highlighted structural features specific to metal activation based on amino acid sequence comparison. Zinc-activated GlxI enzymes such as from human and P. putida possess longer amino acid sequences than that comprising the E. coli enzyme (Figure 6). Additionally, the Saccharomyces cerevisiae and Plasmodium falciparum enzymes appear to be ‘fused dimers’, where a single polypeptide chain assumes the overall tertiary fold observed in the human and E. coli GlxI homodimers [72,125]. In the case of these monomeric, fused, Zn2-activated GlxI sequences, the stretch of polypeptide sequence equivalent to the monomer in the dimeric enzyme is again comparatively longer than the E. coli GlxI amino acid sequence. To determine if the E. coli enzyme’s metal activation profile was unique among GlxI enzymes or whether it represented a novel class of hitherto unrecognized Ni2-activated GlxI, a comparative biochemical study of additional ‘compact’ bacterial GlxI enzymes was pursued. Based on sequence comparisons, several bacterial genomes were scanned for putative GlxI sequences using the E. coli GlxI enzyme as a query [59]. Putative GlxI-encoding genes from Y. pestis, P. aeruginosa and the previously identified, but uncharacterized enzyme from N. meningitidis were cloned into E. coli cell lines for recombinant protein production (Figure 6) [67,79]. All three enzymes were expressed abundantly and possessed GlxI activity. Initial screens showed that substantial quantities of apoenzyme were overproduced in each case, as GlxI activity in crude lysates could be augmented with the addition of Ni2 and Co2, but not Zn2. In parallel Met. Ions Life Sci. 2, 445–472 (2007)


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Figure 6. Sequence alignment for several GlxI enzymes from various sources. Accession numbers (NCBI) for the sequences are included in parentheses after the organism name. Alignment contains GlxI sequences from E. coli (U57363), Y. pestis (CAC91186), Leishmania major (XM_838313), P. aeruginosa (AAG06912), N. meningitidis (CAA74673), P. putida (AAN69360), H. sapiens (AAB49495). Active site residues are marked with an asterisk and loop regions in the longer GlxI sequences are indicated in bold. Alignments were initially obtained through the Clustal W alignment tool, and manually adjusted for gaps in sequences [138,139].

with E. coli GlxI, these additional GlxI sequences could be purified with ease as apoenzymes for metal reconstitution studies. Titration of all three apoenzymes with Ni2, Co2 and Zn2 confirmed preliminary screens that these bacterial GlxI sequences conferred a non-Zn2-dependent Ni2/Co2 activation profile similar to the E. coli enzyme (Table 1). Although the metal activation profiles for these GlxI were similar to the E. coli enzyme, it was observed that the ratio of nickel to cobalt activation did vary with the source of the enzyme; the GlxI from Y. pestis was found to have a substantial enhancement of nickel activation over cobalt activation. Kinetic parameters (Km and Vmax) for these bacterial GlxI enzymes with Ni2 and Co2 reconstitution were within the range observed for the previously characterized E. coli GlxI [79]. Met. Ions Life Sci. 2, 445–472 (2007)

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Table 1. % Maximal activity of bacterial Glyoxalase I enzymes reconstituted with various metal ions. a Metal ion Apo Zn2 Mg2 Ca2 Cd2 Mn2 Co2 Ni2 a

E. coli 1.3 1.3 1.3 1.3 6.8 8.1 30.9 100

P. aeruginosa 0.9 1 1.7 1.7 6.3 6.6 31.5 100

Y. pestis 0 0.4 0.6 0.5 2.4 4.3 23 100

N. meningitidis 0 6.8 0 0 36.7 17.7 72.6 100

Adapted from data reported in [78,79].

Analysis of the sequences for these additional GlxI indicated that all three were in fact shorter than polypeptide segments comprising the zinc-activated P. putida and human GlxI enzymes. Including E. coli GlxI with these newly characterized GlxI enzymes, sequence comparison reveals the presence of positionally equivalent gaps in their amino acid sequence that correspond to short stretches of additional primary structure in the Zn2-activated enzymes (Figure 6). It is tempting to speculate that these additional residues are contributing influences to the metal selectivity of the Zn2-dependent GlxI enzymes and that their absence favors an alternative activation profile as observed for the E. coli enzyme and related orthologs. This structure–function relationship is being actively studied in our laboratory through mutagenesis experiments on the E. coli GlxI enzyme. One obvious attribute for investigation in GlxI structure and function is the variation in active site ligands. Three of the four active site residues in E. coli GlxI are conserved in the human enzyme with the exception of His5, which corresponds to Gln34 in the human active site. The GlxI sequences from Y. pestis, N. meningitidis, P. aeruginosa, P. putida, and Saccharomyces cerevisiae all possess a His residue analogous to His5 in the E. coli enzyme. However, variation of that residue to Gln can be observed in the GlxI sequences from additional multicellular eukaryal GlxI sequences enzymes such as that from soybean [66]. That the His/Gln variation may correlate to differences in metal selectivity is not so straightforward. The P. putida GlxI is a confirmed Zn2-activated enzyme with a His residue analogous to the E. coli enzyme, which does not demonstrate Zn2 activation. Given the structural subtlety of differences that may delineate one GlxI metal activation class from another, mutagenesis was performed on the His5 residue of E. coli GlxI to assess the effect of modifying this variant residue [102]. Gln was substituted for His5 in the E. coli enzyme, generating a variant with a ‘humanized’ active site. Interestingly detectable, but Met. Ions Life Sci. 2, 445–472 (2007)


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low levels of enzyme activation in the presence of Zn2 were now detectable for the variant [102]. Metal reconstitution experiments revealed that the H5Q GlxI was still predominantly Ni2-/Co2-activated, but the metal affinity of the enzyme was severely impaired. Although modification of this active site ligand was influential for producing detectable Zn2-activation, other as yet undefined structural factors appear to be responsible for conferring a robust level of Zn2activation. For the E. coli enzyme it has been shown that increased levels of nickel in the growth medium for cells expressing this recombinant form of the protein resulted in increased levels of nickel incorporation into the enzyme [78]. This was also observed for zinc, but the resulting zinc-bound enzyme was found to be inactive. Although it has been shown that a number of nickel enzymes may require metallochaperones to facilitate nickel incorporation, the exact nature of the nickel incorporation into E. coli’s GlxI is not understood although it is under active investigation.

5.

GLYOXALASE I AS A MEMBER OF THE βαβββ SUPERFAMILY OF PROTEINS

The first reported crystal structure of a GlxI, that of the human enzyme, presented information with respect to structural similarity of the GlxI fold with that of several other proteins in to what has since become known as the βαβββ superfamily of proteins [113]. This is due to the observation that the three-dimensional structure of the GlxI enzyme is a repeat of this motif and that there are a number of proteins that have similar folds. Since the first recognition of this fact by Bergdoll, the list of structurally similar folded proteins has grown to include the following: catechol dioxygenases, mitomycin resistance protein, fosfomycin resistance protein (FosA), and methylmalonyl CoA epimerase [113,126–130]. As observed with many other well-defined structural superfamilies of proteins, sequence conservation between members is quite low in spite of the striking retention of topology. Tandem modules of βαβββ motifs are generally found in the monomeric subunits of these proteins and, the interface between motifs from the same monomer (non-domain-swapped) or different subunits (domain-swapped dimer) form a conserved cleft [131]. This cup-shaped region in βαβββ proteins is functionally conserved as a ligand binding site and also a metal binding site in the case of metalloenzyme representatives of the superfamily. Methylmalonyl CoA epimerase is another metalloenzyme in this superfamily that catalyzes proton transfer using a highly similar active site to GlxI to carry out the R to S conversion of substrate [130]. A crystal structure has been obtained for this enzyme [130], with a metallated active site, but unlike GlxI the active site is contained within one monomer. However, it appears that the topology contained within this enzymatic Met. Ions Life Sci. 2, 445–472 (2007)

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scaffold has been optimized by nature to competently facilitate proton transfer reactions. Of additional interest is another GSH utilizing metalloenzyme in this superfamily, the Mn2-dependent fosfomycin resistance protein FosA from P. aeruginosa. FosA catalyzes the inactivation of the epoxide antibiotic fosfomycin via epoxide ring opening by addition of GSH [132]. The presence of another GSH utilizing enzyme aside from GlxI in this superfamily also suggests that the metal centers in such proteins may be modified through evolution to permit different enzyme activities while maintaining a given co-substrate utilization.

6.

OTHER ASPECTS OF GLYOXALASE I

Although GSH is a widespread intracellular thiol, there are other intracellular thiols of importance [133,134]. If the formation of MG also occurs within these other organisms, it is likely that a GlxI enzyme making use of a non-GSH thiol will be necessary. Interestingly, in the case of the parasite Leishmania major, recent reports have described the isolation and characterization, both enzymological as well as structural of a trypanothione-utilizing GlxI enzyme (Figure 7) [135,136]. The enzyme utilizes the hemithioacetal of MG and trypanothione preferentially. An interesting aspect of this discovery is that L. major GlxI is a Ni2-dependent enzyme. The sequence of this enzyme has been shown to be of

+H N 3 –O

O

H N

H

2C

N H

O O –O

2C

H

N H

NH3+

CH2SH CH2SH H N

H N (CH2)3

O

NH2+ O (CH2)4 N H

O

Trypanothione H3N H

H N

OOC O

O N H CH2SH

H N

H2 N

NH3

O

1-monoglutathionylspermidine

Figure 7. Chemical structures of (A) trypanothione and (B) N1-monoglutathionylspermidine. Met. Ions Life Sci. 2, 445–472 (2007)


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similar size to the shorter E. coli enzyme and displays a remarkably similar metal activation profile [135,136]. Furthermore, the recently published crystal structure for L. major GlxI is quite comparable to the E. coli enzyme. Consistent with the metal binding properties of E. coli GlxI, the L. major enzyme binds one mol of Ni2 per mol dimeric enzyme and interestingly the single metal site occupancy is also apparent in its crystal structure [135,136]. So it appears, at least in this case, that the glyoxalase system is conserved as a route for MG detoxification in cellular environments where GSH is not the predominating intracellular thiol. In addition, it would appear that the non-Zn2-dependent GlxI has evolved an alternative specificity for thiol co-substrates with a marked selectivity for trypanothione over GSH in the case of L. major. This raises interesting questions concerning the substrate specificity of GlxI enzymes in relation to metal dependence. Considering the substantial intracellular pool of glutathionylspermidine in E. coli under stationary or anaerobic growth conditions [137], further characterization of its Ni2-activated GlxI may provide insight as to whether E. coli GlxI can accept larger, more complex thiol co-substrates, mirroring the preference for larger thiols observed in the leishmanial enzyme. The systematic differences in length and secondary structural elements between the two metal activation classes of GlxI must be examined to uncover the structural origin of differential metal specificities. If these determinants also correlate to substrate/co-substrate utilization profiles in one class versus another, these features may be exploited in the development of inhibitors targeting bacterial and parasitic GlxI enzymes.

7.

CONCLUSIONS

In spite of the importance of the detoxification/control of MG levels in cells, Nature has interestingly evolved two highly related GlxI enzymes which possess very different metal activation profiles. Although nickel is frequently detected as a redox active metal in enzymes such as hydrogenase, methyl CoM reductase, and others, GlxI, like urease, is another metal activated non-redox enzyme that exhibits Ni2-selectivity [95,100]. Given the prevalence of Zn2-dependent GlxI enzymes and the suitability of Zn2 to accommodate changes in coordination environment during the catalytic cycle the presence of Ni2-dependent/non-Zn2-activated GlxI is remarkable. The role of Ni2 in GlxI-catalyzed isomerization is an area of research that can serve as a basis to address interesting questions concerning how protein structure might influence selectivity for a particular metal ion versus another for the same catalyzed reaction. Given the prevalence of non-Zn2-dependent GlxI enzymes in prokaryotic organisms, metal selectivity of this enzyme may have diverged in parallel to the establishment of eukaryotic and prokaryotic lineages. Certainly, the structural differences between Zn2-dependent and non-Zn2-dependent Met. Ions Life Sci. 2, 445–472 (2007)

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GlxI, may correlate to coordination environments better suited for one type of metal activation versus another.

ACKNOWLEDGMENTS The authors wish to thank Dr J. F. J. Barnard, Dr S. L. Clugston, R. Kinach, H. D. Ly, D. Miedema, and R. Yajima for previous contributions to various parts of the research described in this article and to gratefully acknowledge our collaborators, Dr G. Davidson, Dr M. M. He, Dr D. Markham, Dr M. J. Maroney, and Dr B. W. Matthews. The authors would also like to thank NSERC (Canada) for a Discovery Grant (JFH) and a graduate scholarship (NS) and the University of Waterloo for additional support.

ABBREVIATIONS AGE CoA CoM DHAP DSC EDTA EXAFS FBP FosA GlxI GlxII HIPC-GSH GSH ICP-AES ITC LB MG NADPH NBC-GSH NCBI Pi SDS-PAGE TIM XANES XAS

advanced glycation end product coenzyme A coenzyme M dihydroxyacetone phosphate differential scanning calorimetry ethylenediamine-N,N,N,N-tetraacetate extended X-ray absorption fine structure fructose-1,6-bisphosphate aldolase fosfomycin resistance protein glyoxalase I ( S-D-lactoylglutathione methylglyoxal lyase) glyoxalase II ( S-2-hydroxyacylglutathione hydrolase) S-(N-hydroxy-N-p-iodophenylcarbamoyl)glutathione glutathione inductively coupled plasma atomic emission spectroscopy isothermal titration calorimetry Luria Bertani medium methylglyoxal nicotinamide adenine dinucleotide phosphate, reduced S-p-nitrobenzyloxycarbonylglutathione National Center for Biotechnology Information inorganic phosphate sodium dodecylsulfate polyacrylamide gel electrophoresis triose phosphate isomerase X-ray absorption near-edge structure X-ray absorption spectroscopy

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12 Nickel in Acireductone Dioxygenase Thomas C. Pochapsky, Tingting Ju, Marina Dang, Rachel Beaulieu, Gina M. Pagani, and Bo OuYang Departments of Chemistry and Biochemistry, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454-9110, USA <[email protected]>

1. INTRODUCTION 2. THE METHIONINE SALVAGE PATHWAY 3. ONE PROTEIN, TWO ENZYMES: ACIREDUCTONE DIOXYGENASE FROM Klebsiella oxytoca 4. HOMOLOGS OF ACIREDUCTONE DIOXYGENASE FROM OTHER ORGANISMS 5. KNOWN ACIREDUCTONE DIOXYGENASE STRUCTURES 6. SPECTROSCOPIC PROBES OF ACIREDUCTONE DIOXYGENASE ENZYME ACTIVE SITES 7. ENZYMATIC STUDIES OF ACIREDUCTONE DIOXYGENASE 8. MECHANISTIC CONSIDERATIONS: WHAT IS THE ROLE OF Ni(II) IN ACIREDUCTONE DIOXYGENASE ACTIVITY? 9. STRUCTURALLY AND FUNCTIONALLY RELATED ENZYMES 10. FUTURE DIRECTIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES



474 475 477 481 483 486 489 490 493 495 497 497 498

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Nickel plays a role in a variety of biological processes, including redox-active roles in hydrogen and carbon monoxide metabolism in microorganisms under anaerobic conditions [1]. However, to date, no mammalian enzyme has been positively found to contain nickel in vivo, and the importance of Ni as a trace element in human nutrition is not well documented. Therefore, the discovery of a nickel-dependent bacterial enzyme in the ubiquitous methionine salvage pathway, or MSP (see Figure 1 below), has led to speculation that a mammalian nickel-dependent enzyme might yet be discovered. The nickel-dependent bacterial enzyme is acireductone dioxygenase (ARD) from the bacterium Klebsiella oxytoca (formerly K. pneumoniae) ATCC strain 8724 [2–5], and the nickel-containing form of the enzyme catalyzes an off-pathway shunt from the MSP that results in the formation of relatively large amounts of carbon monoxide by K. oxytoca under normal growth conditions [3]. More remarkably, it was found that over-expression of the ARD gene in E. coli yields two forms of ARD, one that catalyzes the off-pathway shunt reaction, and one that catalyzes the on-pathway reaction leading to the formation of methionine [4]. The two enzymes are chromatographically separable on ion exchange and hydrophobic interaction phases, but are formed from identical polypeptides, with the only chemical difference between them being the identity of the bound metal. The on-pathway enzyme (originally called E2′ and now called ARD′) binds approximately one equivalent of Fe2, while the off-pathway shunt enzyme (originally E2 and now ARD) contains one equivalent of Ni2 per mole of polypeptide [5]. As isolated from E. coli, both enzymes are monomers, suggesting that their chromatographic separability is due to differences in structure rather than oligomerization state. It was demonstrated that metal ion removal and replacement yielded the appropriate activity, regardless of which form of the enzyme was used; that is, reconstitution of either form of the enzyme with NiCl2 yields ARD activity, while reconstitution with FeCl2 yields ARD′ activity. It was also reported that the enzyme is somewhat promiscuous, and that Ni-type activity is obtained by reconstitution with Co2 and Mn2 and partial Fe-type activity by reconstitution with Mg2 [6]. ARD shows tighter binding of Ni2 than Fe2, as demonstrated by the fact that Fe can be removed from the folded enzyme by dialysis against EDTA, while Ni removal requires that the protein be unfolded [4]. X-ray absorption spectroscopy (XAS) data, including XANES (X-ray absorption near-edge spectroscopy) and EXAFS (extended X-ray absorption fine structure), are consistent with octahedral coordination geometry of Ni2 in ARD by N/O with three histidine ligands [7]. From NMR structural studies and sequence similarity (see Sections 4 and 5), the protein-based ligands for metal ions in ARD are proposed to be His 96, His 98, His 140 and Glu 102 [8]. These residues are strictly conserved within ARD-type genes identified in multiple genomes. The proposed ligation scheme is consistent with the crystal structure of an ARD homologue from Mus musculus Met. Ions Life Sci. 2, 473–500 (2007)

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(house mouse) (PDB accession 1VR3), although the identity of the bound metal in that structure is unknown. This proposal is also supported by a mutagenesis study on the ARD from yeast, that showed Glu 91 (the residue corresponding to Glu 102 in K. oxytoca ARD) is essential for function [9].

2.

THE METHIONINE SALVAGE PATHWAY

The amino acid methionine is an essential amino acid in human nutrition and, apart from its role in protein structure and biosynthesis, is an important intermediate in biosynthetic pathways requiring alkyl carbons activated towards nucleophiles (Figure 1). S-adenosylmethionine (SAM or Ado-Met) 1 is formed enzymatically from ATP and methionine, and contains a cationic sulfonium species that activates all three carbons bonded to the sulfur. The activated sulfonium methyl of SAM is used by a variety of methyl transferases (e.g., for methylation of DNA and ethanolamine in complex lipids). The removal of the sulfonium methyl from SAM results in formation of S-adenosylhomocysteine, which in turn is

Figure 1. The methionine salvage pathway. Enzyme abbreviations are shown in bold and refer to the discussion in the text. Met. Ions Life Sci. 2, 473–500 (2007)

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COO– +

O

S

H 3N

H

H

H

OH

H OH

O

S-adenosylmethionine (SAM) 1

+H

O–

3N +

SAM decarboxylase

H3N

ornithine

CO2

ODC Ade

COO– +H

3N

+

H3N

O

S H

H

H

OH

H OH

+H

NH3+

3N

putrescine

ethylene (in plants)

+

H3N

Ade

N H2+

NH3+

spermidine

S O H

H

H

OH

H OH

Methylthioadenosine (MTA) 2

+H

H2+ N 3N

N H2+

NH3+

spermine

Figure 2. The role of SAM in polyamine and ethylene biosyntheses leading to the formation of MTA 2. See text for a more complete discussion.

remethylated to methionione by methylcobalamin. A more complete discussion of this pathway can be found in [10]. Another important role of SAM is in the synthesis of the polyamines spermine and spermidine (Figure 2). While the physiological roles of polyamines are as yet unclear, upregulation of their biosynthesis is associated with the cell cycle and cell growth, including suppression of apoptosis in developing tissues, while their synthesis is downregulated in senescent cells and mature tissue [11]. Defects in polyamine regulation are associated with oncogenesis, and neoplastic cells from colorectal cancers are found to have higher levels of polyamines than normal tissue [12]. The first step in polyamine biosynthesis is the decarboxylation of the amino acid ornithine by ornithine decarboxylase (ODC), yielding putrescine (1,4diaminobutane). Successive aminopropylation of the amino groups of putrescine yields spermidine and spermine, respectively. The aminopropyl groups required for spermidine and spermine are provided by SAM after decarboxylation of the methionine moiety, a reaction catalyzed by SAM decarboxylase. The biproduct Met. Ions Life Sci. 2, 473–500 (2007)


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of aminopropyl transfer is methylthioadenosine (MTA) 2, a metabolite that has a strong regulatory affect on polyamine synthesis and retention [13] and on transmethylation [14,15]. MTA has been implicated in the inflammatory response [16,17] and exhibits both pro- and antiapoptotic effector activity, depending upon concentration and context [18]. Due to the regulatory roles played by MTA, intracellular levels of this compound are quite low under normal conditions (3 nmol/g in rat organs) [19]. The methionine salvage pathway is critical in maintaining MTA homeostasis, and makes use of the carbon skeleton of the ribose moiety of MTA to regenerate methionine, salvaging the thiomethyl group of the original methionine molecule. The first steps in the MSP are the deadenylation and phosphorylation to form methylthioribose-1-phosphate 3 (Fig. 1) and adenine. In eukaryotes and some prokaryotes, these reactions are catalyzed by a single enzyme, MTA phosphorylase (MTAP) [20,21]. However, in many bacteria, including K. oxytoca, there are separate enzymes for each reaction, MTA nucleosidase (MTAN) and 5methylthioribose kinase (MTRK) [21-25]. Methylthioribose-1-phosphate is then converted to methylthioribulose-1-phosphate 4 (Figure 1) by methylthioribose-1phosphate isomerase (MTR-1-PI) [26]. Compound 4 is then dehydrated (compound 5) and tautomerized to yield 1-phosphonooxy-2,3-diketo-5-thiomethylpentane 6 (Figure 1). Although the dehydratase responsible for the transformation of 4 to 6 has not been identified in K. oxytoca, a gene product responsible for this chemistry has been identified in B. subtilis [21]. Based on sequence homology, the dehydratase is a member of the class II aldolase family (Pfam0096, pfam. wustl.edu) and most genomes, including K. oxytoca, have a corresponding open reading frame associated with genes from the methionine salvage pathway. Under acidic conditions, des-thio analogs of 6 form the stable hydrate 6a [27,28]. In K. oxytoca, compound 6 is the substrate for E1 enolase/phosphatase, a member of the haloacid dehalogenase superfamily [27–30], and a corresponding protein has been found in H. sapiens [31]. This enzyme catalyzes the formation of 2-hydroxy-3-keto-5-thiomethylpent-1-ene 7 (Figure 1) from compound 6. Interestingly, in some organisms (e.g., B. subtilis) the enolase reaction is catalyzed by a RuBisCO (ribulose-1,5-bisphosphate carboxylase/oxygenase) homolog [32] and the phosphatase reaction is catalyzed by a separate gene product [21].

3.

ONE PROTEIN, TWO ENZYMES: ACIREDUCTONE DIOXYGENASE FROM Klebsiella oxytoca

Compound 7 is often called by the generic name acireductone or aci-reductone, referring to the 1,2-dihydroxy-1-ene-3-carbonyl functionality, also found in ascorbic acid. Acireductone 7 is the pentultimate intermediate in the MSP, and represents a branch-point in the MSP. The molecule is quite reactive towards Met. Ions Life Sci. 2, 473–500 (2007)

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OH OH

7 O OH OH

7a O OH OH

7b

oxidizing agents, and homologs of 7 are sufficiently acidic that they are singly deprotonated at physiological pH in aqueous solution [5]. Furthermore, the reaction with molecular oxygen is sufficiently fast under non-enzymatic conditions that it is inconvenient to store acireductones of type 7 for any extended period of time; acireductone for enzymatic assays is typically generated in situ by treatment of the diketophosphonate precursor with enolase/phosphatase immediately prior to the assay. The laboratory synthesis of precursor compound 6 (Figure 1) has not been reported, but a convenient synthesis of des-thio analogs of 6 (precursors for compounds 7a and 7b), suitable for MSP mechanistic and activity assays has been described [28]. Because of acireductone’s reactivity towards molecular oxygen, it was originally assumed that the pentultimate step in the MSP, the oxidative cleavage of 7 by dioxygen to form 2-keto-4-(methylthio)butyrate 8 (Figure 1), and formate, was non-enzymatic [27]. However, studies of the metabolic fate of methylthioribose-1-phosphate 3 in the methionine salvage pathway of K. oxytoca indicated that a substantial fraction of 3 is diverted to the formation of methylthiopropionic acid 9 (Figure 1) [26]. Furthermore, it was shown that keto-acid 8 was not the source of 9, but that the branch point occurred earlier in the pathway [26]. The search for the off-pathway shunt to compound 9 led to the identification of an enzyme fraction that catalyzed the formation of butyric acid, formate, and carbon monoxide from acireductone 7 [3]. CO-producing enzymes are rare in Met. Ions Life Sci. 2, 473–500 (2007)


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nature, and the role of this enzyme product in K. oxytoca remains unclear, although this reaction occurs in the wild-type organism [26]. Experiments with rat liver homogenates identified the on-pathway MSP reaction product 8 resulting from the metabolism of MTA, but no evidence for carbon monoxide formation [3]. Still, K. oxytoca is not unique in the production of CO via this mechanism, as CO production from methylthioribose has been reported for B. subtilis [21]. The CO-producing enzyme, designated E2, was purified and characterized, and found to be an 18.5 kDa monomer [3]. No cofactor or bound metal was initially identified, although a Mg2 dependence was suggested. This dependence was in fact later found to be due to the enolase–phosphatase enzyme required to generate acireductone from precursor [27]. Upon cloning and over-expression in E. coli, the surprising observation was made that two active protein fractions were obtained. One fraction catalyzed the off-pathway shunt from 7 to 9 plus CO and formate, while the other fraction catalyzed the on-pathway reaction leading from 7 to 8 plus formate [4]. The two fractions were separable by ion exchange and hydrophobic interaction chromatographies, suggesting significant structural differences between them. However, both proteins were monomeric, and the only chemical difference between them was the nature of the metal ion bound in each case: the off-pathway enzyme (E2, later named ARD) bound one equivalent of Ni2, while the on-pathway form (E2′, later named ARD′) contained one equivalent of Fe2. Removal of the metal from either form of the enzyme, followed by reconstitution of the apo-protein with Ni2 gave the off-pathway reaction, while reconstitution with Fe2 of apo-protein generated from either form gave the onpathway chemistry. It was observed that other divalent metals could be used to reconstitute activity, with Co2 providing a good substitute for Ni2 in producing ARD (off-pathway) activity [4]. Indeed, metal analysis of ARD isolated from K. oxytoca indicates that about 20% of the enzyme contains cobalt rather than nickel [5]. It was also found that both Mn2 and Co2 could replace Ni2 in the enzyme, giving ARD activity, while Mg2 appeared to give partial ARD′ activity [6]. Addition of apo-ARD to an equimolar mixture of Fe2 and Ni2 produced 80% of the Ni-containing form within one minute, indicating a substantial preference for Ni2 binding over Fe2 [4]. The promiscuity of K. oxytoca ARD (KoARD) toward different metals as well as the activity of these different forms suggests that the redox properties of the bound metal are relatively unimportant for the chemistry catalyzed by ARD. Enzyme–substrate complexes of both the Fe-containing and Ni-containing forms of KoARD are EPR-silent under both aerobic and anaerobic conditions, suggesting that if any metal-based redox chemistry occurs in either enzyme, it is transitory [5]. Furthermore, there is no long-lived organic radical intermediate in the absence of oxygen, although a cyclopropyl substrate analog 7b that gives rise to a reactive radical intermediate does slowly inactivate both enzymes [5]. This would suggest that either a transient radical intermediate is formed in the ternary O2-7-enzyme complex or that a radical-based side reaction can occur. Finally, O2 Met. Ions Life Sci. 2, 473–500 (2007)

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is not bound by either enzyme in the absence of substrate, indicating that a direct O2-metal bond is unlikely [5]. Based on the observation of a spectral shift in the substrate upon binding stoichiometrically to ARD and ARD′, it was concluded that the substrate bound as a dianion [5]. While this is suggestive of a direct metal–substrate interaction, the first direct observations concerning the interactions between metal and substrate in Ni-containing ARD were made using 1H NMR and XAS [7]. From hyperfine shift patterns in the downfield region of the 1H NMR spectrum of Ni-containing ARD, it was concluded that three His residues ligate the Ni2 ion, two through the Nε nitrogen and one through the Nδ. Addition of substrate under anaerobic conditions results in complex perturbations of the His ligand resonances, indicating a direct interaction between the metal and substrate in the enzyme-substrate (ES) complex, probably ligation or chelation [7]. Ni EXAFS analysis of ARD and the ARD–7a complex gives further information regarding the metal binding site. The Ni EXAFS data for ARD is consistent with octahedral ligation of the Ni2 by O/N ligands, and second-sphere scattering is best fit by 3–4 His ligands. Interestingly, upon addition of substrate, the EXAFS data is best fit by octahedral ligation including 2–3 His ligands, suggesting that one His ligand is displaced by substrate [7]. Gaps in NMR sequential resonance assignments identified four His residues, His 96, His 98, His 137 and His 140 that are paramagnetically broadened, and thus were considered as candidates for ligating the Ni2 [33]. As more members of the ARD family were identified in other organisms, it became clear that His 137 is not conserved, but the other three His residues are strictly conserved (see Figure 3 below). Furthermore, the same patterns of paramagnetic broadening are seen in Fe2-containing ARD′, indicating that Fe2 is bound in the same site as Ni2 in ARD (T. Ju, personal communication), and with similar geometry (M. Maroney, personal communication). Mutation of His 96 and His 140 to Ala result in inactivation and loss of solubility of ARD, while mutation at His 137 does not affect activity [34]. The H98A mutation results in a relatively insoluble and inactive protein; however, replacement of His 98 with Ser results in a soluble well-folded protein, that while not spontaneously binding Ni2, is capable of accepting Ni2 under forcing conditions, resulting in an NMR spectrum similar to the Ni-bound form of ARD. The Ni-bound H98S mutant is active, catalyzing the oxidative formation of CO from 7a [34]. The H98S mutation also affects iron binding. Based on isothermal calorimetry measurements, the affinity of the mutant for either Fe2 or Ni2 is several orders of magnitude below that of wild-type ARD [34]. Besides the conserved histidine residues, there are a number of conserved acidic residues near the metal binding site, including the strictly conserved Glu 95 and Glu 102 and conservatively acidic residues Glu 100 and Asp 101. In this case, structural homology rather than sequence homology identified Glu 102 as a likely fourth ligand for the metal in ARD (see Section 5). The role of the other Met. Ions Life Sci. 2, 473–500 (2007)


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conserved acidic residues is not clear. The side chain of the residue homologous with Glu 95 residue is surface-exposed in the MmARD structure and in both the modeled ARD and experimental H98S ARD structures (see Section 5). Asp 101 forms a salt bridge with Lys 68 in the refined ARD structure, and may provide a structural link between the active site and nearby structural features involved in substrate access and orientation within the active site.

4.

HOMOLOGS OF ACIREDUCTONE DIOXYGENASE FROM OTHER ORGANISMS

Although the first ARD enzyme was discovered relatively recently, it is now seen that most prokaryotic and eukaryotic genomes contain at least one open reading frame that encodes a protein homologous to ARD, and the ARD family has been recognized as a distinct class of enzymes (see pfam.wustl.edu entry PF03079). To date, the best characterized members of the ARD family besides Klebsiella oxytoca ARD include OsARD1 from Oryza sativa (rice) [35], ADI1 from yeast (Saccharomyces cerevisiae) [36], ALP (ARD-like protein) encoded by an androgen-responsive gene from rat (Rattus norvegicus) [37], and MTCPB-1 (membrane-type 1 matrix metalloproteinase cytoplasmic tail binding protein-1, accession BC001467) from Homo sapiens [9,38]. A truncated form of MTCBP1 has been implicated in the replication of hepatitis C virus in otherwise nonpermissive cell lines [39]. A crystal structure for the ARD homolog from house mouse has been deposited in the PDB database (accession number 1VR3) from the Joint Center for Structural Genomics high-throughput structural genomics project [40]. Figure 3 provides a ClustalW alignment of these sequences. Those ARD homologs for which physiological roles have been found all appear to function in the MSP, although they may play regulatory roles in their native organisms as well. A recently published study of the OsARD1 gene in deep-water rice plants indicate that the expression of this gene is strongly upregulated as an early response to ethylene production under hypoxic conditions that result from submergence [35]. Ethylene is biosynthesized from cyclopropylglycine, that in turn is derived from S-adenosylmethionine with concomitant production of MTA, suggesting that the gene product OsARD1 indeed plays a role in the MSP of rice (see Figure 2). OsARD1 (previously known as Sip2, for submergenceinduced protein 2) is catalytically active towards acireductone, although only the Fe-containing form that catalyzes the on-pathway reaction has been isolated from bacterial expression systems. Furthermore, Fe2 in OsARD1 is relatively labile, and purification buffers require ferrous ammonium sulfate and cysteine to be present in order to maintain the iron-bound form of OsARD1. Attempts to reconstitute OsARD1 with Ni2 resulted in an oligomeric species of high molecular weight. Although the Ni-reconstituted OsARD1 catalyzes the offpathway oxidation of 7a with formation of CO, the catalytic activity is lower Met. Ions Life Sci. 2, 473–500 (2007)

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POCHAPSKY et al. ---------------------------MVQAWYMD-DAPGDPRQPHRPDP ----------------------------------------------------------------XGSDKIHHHHHHXVQAWYXD-ESTADPRKPHRAQP ----------------MENEFQDGKTEVIEAWYMD-DSEEDQRLPHHREP ---------------------------MVKVYIHDNKVDSDYRAPHN--S ---------------------SALTIFSVKDPQNSLWHSTNAEEIQQQLN MQCPTGLEEPDGKADTEECPMSSLSVYHVSSPEIPNKVLTHFEDIASTLA -------------------------MATIRIHDEANTTIENQEEVASFLD

GRPVGLEQLRRLGVLYWK--------LDADKYEND--PELEKIRRERNYS -------------------------------------------------DRPVSLEQLRTLGVLYWK--------LDADKYEND--PELEKIRKXRNYS KEFIPVDKLTELGVISWR--------LNPDNWENC--ENLKRIREARGYS GTELSLDELAKLGVIY-------------KYCANE--EEVNEIARQREYK AKGVRFERWQADRDLGAA--------PTAETVIAAYQHAIDKLVAEKGYQ EQGVRFDRWQAAAKIQPG--------ASQEEVIGAYKEQIDKLMTERGYI SQEVIYEQWDITRLPEHLSEKYDLTEEEKQQILDTFETEIKDISTRRGYK . . . .

22 34 33 21 29 50 25

62 74 73 56 71 92 75

WMDIITICKDKLP---NYEEKIKMFYEEHLHLDDEIRYILDGSGYFDVRD -MDIITICKDKLP---NYEEKIKMFYEEHLHLDDEIRYILDGSGYFDVRD WXDIITICKDTLP---NYEEKIKXFFEEHLHLDEEIRYILEGSGYFDVRD YVDICDVCPEKLP---NYETKIKSFFEEHLHTDEEIRYCLEGSGYFDVRD NRDVVNICEGSFKSEAEFNEKLATFYQEHLHEDEEIRYCLEGAGYFDVRD SWDVISLRADNPQ----KEALREKFLNEHTHGEDEVRFFVEGAGLFCLHI TVDVISLNSDHPQ----KAELRAKFLEEHRHGEDEVRFFVAGRGLFTLHI AQDVISLSDSNPK----LDELLENFKREHHHTDDEVRFIVSGHGIFVIQG *: : * .** * ::*:*: : * * * ::

109 46 121 120 106 117 138 121

KE--DQWIRIFMEKGDMVTLPAGIYHRFTVDEKNYTKAMRLFVGEPVWTA KE--DQWIRIFMEKGDMVTLPAGIYHRFTVDEKNYTKAMRLFVGEPVWTA KE--DKWIRISXEKGDXITLPAGIYHRFTLDEKNYVKAXRLFVGEPVWTP QN--DQWIRIALKKGGMIVLPAGMYHRFTLDTDNYIKAMRLFVGDPVWTP ASTPENWIRCLVESGDLLILPPGIYHRFTLTTSNHIKALRLFKDEPKWQA GD---EVFQVLCEKNDLISVPAHTPHWFDMGSEPNFTAIRIFDNPEGWIA DD---YVYAVLCEKNDLISVPAGTKHWFDMGENPHFVAIRLFNNPEGWVA QD--GTFFDVRLNPGDLISVPENIRHYFTLQEDRKVVAVRIFVTTEGWVP . : .. : :* * * : . * *:* * .

157 94 169 168 156 164 185 169

YNRP--ADHFEARGQYVKFLAQTA--------YNRP--ADHFEARGQYVKFLAQTA--------YNRP--ADHFDARVQYXSFLEGTA--------YNRP--HDHLPARKEFLAKLLKSEGENQAVEGF INRSNQADSLPVRKDYIALINQY---------QFTG---DDIASAYPRLA--------------NFTG---EDIAGRFPRLED-------------IYEK---DSVNQ--------------------: .

179 116 191 199 179 179 201 178

Figure 3. Sequence identity and similarity between ARD homologues. Asterisks (*) indicate complete conservation, colons (:) indicate conservative replacement in the row underneath sequence alignments. Conserved residues proposed to be ligands for metal are indicated by an arrow above the alignment. Alignment was performed using the ClustalW algorithm [67].

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than that of the on-pathway enzyme Fe-OsARD1 and much lower than that of the bacterial Ni-containing ARD [35]. This suggests that the production of CO by this enzyme is unlikely to be important physiologically. A second ARD homolog, OsARD2, is expressed at low levels in multiple tissues in rice, but the gene product has not been characterized [35]. The human homolog of ARD was first detected as a cDNA from human liver, the gene product of which enables hepatitis C virus to replicate in an otherwise non-permissive cell line [39]. This gene encoded an N-terminal truncation of longer ORFs identified from human ovarian cancer and choriocarcinoma cell lines with high homology to the Sip2 rice gene product, later identified as OsARD1. This liver cDNA gene product was therefore termed SipL (for submergence-induced protein-like protein) [39]. More recently, the fulllength 179-residue human ARD homolog was identified as a regulator of matrix metalloproteinase activity [38]. It was found that the full-length gene product (termed MTCBP-1) associated with the cytoplasmic tail of membrane-type 1 matrix metalloproteinase MT1-MMP/MMP-14, and suppressed the activity of that proteinase in cell migration and tissue invasion. Interestingly, expression of MTCBP-1 was lower in human tumor cell lines than in non-transformed fibroblasts, suggesting a link between metastasis (a process that requires active matrix metalloproteinase) and expression of MTCBP-1 [38]. The same group that published the initial report concerning MTCBP-1 later showed that this gene product restored the ability of yeast cells in which the yeast homolog of the ARD gene had been knocked out (YMR009w) to grow in sulfur-depleted media supplemented with MTA [9]. These results indicate that MTCBP-1 acts as an acireductone dioxygenase in vivo in eukaryotes. The rat homolog of ARD, ALP1 (ARD-like-protein 1) is closely related to MTCBP-1, and was observed to be upregulated by androgen treatment in rat prostate, a common therapy for prostate tumors [37].

5.

KNOWN ACIREDUCTONE DIOXYGENASE STRUCTURES

ARD family proteins belong to a functionally diverse structural superfamily known as cupins [41,42]. Cupins are characterized by a conserved antiparallel β -helix that gives rise to a classic ‘jelly-roll’ motif, and include a wide range of proteins, both enzymic and non-enzymic in nature. There are currently three ARD structures available in the RCSB PDB, the original NMR-determined model of NiARD from Klebsiella (Ni-KoARD, PDB entry 1M40) [8], the crystallographic structure of the mouse homolog MmARD (1VR3) [40] and a recently published refinement of the Ni-KoARD structure incorporating residual dipolar couplings (1ZRR) [43] (Figure 4). The original structure of Ni-KoARD confirmed the presence of the cupin fold, and presented a model for active site Ni binding based on XAS, paramagnetic NMR spectroscopy and sequence homology [7]. Met. Ions Life Sci. 2, 473–500 (2007)

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Figure 4. (A) Ribbon structure of NiARD (PDB entry 1ZRR). Letters reference to the ARD sequence as follows: A (Ala 2-Phe 6), B (Leu 15-Ser 18), C (Glu 23-Lys 31), D (Val 33-Glu 36), E (Thr 52-Tyr 57), E′ (Ile 61-Lys 68), F (Ser 72-Leu 78), G (Lys 85-Glu 90), H (Phe 92-Glu 95), I (Arg 104-Val 107), J (Gly 111-Ile 117), K (Glu 120-Leu 125), L (Asn 129Ile 132), M (His 140-Met 144), N (Phe 150-Phe 156), O (Trp 162-Phe 166), P (Ile 171-Ala 174). Position of Ni2 is shown as a blue sphere. (B) Crystallographic structure of MmARD from Mus musculus (PDB entry 1VR3 [40]) with structural features homologous to NiARD indicated by lettering. Structure is viewed in approximately the same orientation as NiARD in part (A). The same letters are used to indicate structurally homologous features. A (Val 2-Tyr 6), B (Arg 24-Ser 27), C (Glu 29-Leu 34), D (Val 35-Trp 39), E (Lys 45-Asn 48), E′ (Leu 52-Arg 59), F (Ser 62-Ile 69), G (Asn 76-Lys 82), H (Phe 84-Glu 87), I (Arg 96-Leu 99), J (Gly 103-Asp 109), K (Lys 113-Ser 118), L (Gly 122-Ile 125), M (His 133-Thr 136), N (Val 143-Phe 149), O (Trp 155-Asn 159) and P (Gln 170-Ala 179). Position of metal (presumed to be Ni2) is shown as blue sphere. (C) Heavy atom positions of the refined NiARD solution structure. Paramagnetically affected residues are shown in bold. Color key: red, Trp 162-Phe 166; purple, Asn 94-Val 103; yellow, Val 134-Asp 143. Position of Ni2 indicated by a green sphere. (D) Metal binding site of NiARD based on modeling as described in the text. Position of Ni2 indicated by a green sphere. View is from the opening to the active site (over strand N, with helix G to the upper left). Reproduced by kind permission of Springer Science and Business Media from [43]. Figures generated by MOLMOL [68].


485

Due to the paramagnetism of the bound nickel ion, standard proton-detected NMR experiments cannot be used to provide direct structural information within ⬃9 Å of the metal, and so structural details of the active site could not be determined for the original structure. However, the paramagnetic broadening patterns observed in the NMR spectra of NiARD confirm those parts of the protein that are sequentially and/or spatially adjacent to the active site; these include residues 95-103, 135-144 and 162-166. The conserved His residues 96, 98 and 140 all lie in this region, as do a number of conserved or conservatively substituted acidic residues. At the time of the original publication, there was no closely homologous structure, so a metal ligation scheme was proposed based on the structure of jack bean canavalin [44], a cupin with a ‘knuckle’ of amino acid side chains that provided a scaffold into which the Ni-binding site of ARD could be modeled [8]. The resulting ligation scheme, shown in Figure 4d, provides two open cis equatorial ligand sites which would presumably be the site of substrate binding. The recently deposited MmARD structure supports this ligation scheme (see Figures 4b and 9 below). As proposed for ARD, the two non-protein ligands are equatorial and cis with respect to each other. However, the nature of the two equatorial non-protein ligands in the MmARD structure are unknown; they do not appear to be water. Furthermore, the metal bound in the MmARD active site is unknown, and the activity of MmARD towards acireductone has not been reported. However, the closely related human homolog HsARD does exhibit ARD activity when overexpressed in bacterial cells (G. Pagani and M. Dang, unpublished), so it is likely that the MmARD structure is that of an active enzyme. The MmARD structure also provides insight into the position of residues 162–166 in the NiARD structure. In the original calculations of the 1M40 structure, no restraints were available other than paramagnetic broadening for residues 162–166, and they appear in the 1M40 structure as an unrestrained segment polypeptide behind the active site. However, the MmARD structure indicates that a short antiparallel hydrogen bonding arrangement exists between what would be residue 97 in NiARD and Ile 163 in this short stretch of polypeptide. This places the Trp 162 indole in a position homologous with that of Trp 155 in MmARD, partially occluding access to the active site and likely enforcing a steric requirement on the ligation geometry of bound ligand (see Figure 4d). The 1ZRR structure incorporates this hydrogen bond as a model, which results in a significant repositioning of the polypeptide immediately preceding Trp 162, as well as the C-terminal end of helix E′, that interacts with Phe 156 and Asn 158. Despite relatively low sequence homology (23% identity, 41% similarity), MmARD (1VR3) and KoARD (1ZRR) show considerable structural conservation (see Figure 4a and b). Corresponding secondary structural features can be identified in both structures with the exception of the C-terminal residues. Unlike the C-terminal 3,10-helix (residues 171–175) that forms a perpendicular cap across the top of the β -barrel in Ni-KoARD, the C-terminal polypeptide of MmARD forms a long helix the axis of which lies parallel to the long axis of the β -barrel. Met. Ions Life Sci. 2, 473–500 (2007)

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Instead of the helical cap, the top of the β -barrel in MmARD is covered by a long section of polypeptide inserted between the first two N-terminal strands of the β -barrel for which there is no homologous peptide in the KoARD sequence. It is at present unclear whether this difference is significant, given the apparent similarity between the active sites of the two structures. However, preliminary structural data for the Fe-containing form of KoARD (ARD′) suggests that the C-terminal of this form is unstructured and does not form the cross-barrel cap (T. Ju, unpublished). If this is the case, it is not clear which form of the enzyme, ARD or ARD′, the MmARD structure represents. One of the most surprising aspects of the ‘one protein, two enzymes’ story is the observation of different physical properties for ARD and ARD′, particularly chromatographic separability on ion exchange and hydrophobic columns. These differences imply significant structural differences between the two enzymes. However, the nature of these differences, other than the absence of the C-terminal cap in ARD′, are currently unclear. The possibility that the differences are due to different oligomerization states for the two proteins appears to be ruled out by the retention times of both enzymes on size exclusion chromatography media, which correspond to monomers of the correct molecular weight [5]. We note that a different situation may exist with eukaryotic ARDs; while the mouse homolog appears to be monomeric, the rice-derived OsARD1 is trimeric [35].

6. SPECTROSCOPIC PROBES OF ACIREDUCTONE DIOXYGENASE ENZYME ACTIVE SITES Ni2 has a d8 electronic configuration, and with pure octahedral geometry by O/N ligands or weak tetrahedral distortion is expected to have S 1, rendering it EPRsilent in the perpendicular EPR mode. Although weak electronic transitions at 25 000, 14 000 and 9 000 cm1 are expected [45], no UV/visible absorption bands other than aromatic amino acid bands are detected for NiARD at concentrations lower than 1 mM. To date, the most reliable spectroscopic probe of the Ni-bound ARD active site is X-ray absorption spectroscopy. XANES spectra for both the resting-state and enzyme–substrate complex of NiARD are consistent with octahedral N/O ligation geometry, with a peak observed at 8332 eV [7]. The lack of a peak or shoulder near 8338 eV effectively rules out square planar or square pyramidal geometry for either the resting state or ES forms (Figure 5). Furthermore, a small shift of the near-edge absorption (0.4 eV) upon substrate binding indicates that the oxidation state of the Ni does not change upon substrate binding. Fourier analysis of EXAFS spectra are consistent with three or four His ligands of Ni2 in the resting state of ARD, while the EXAFS spectrum of the ES complex suggests that one of the His residues is displaced by substrate binding [7]. Based on structural considerations, only His 98 is located in a Met. Ions Life Sci. 2, 473–500 (2007)

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Figure 5. Top: Ni K-edge XANES of the resting (top) and the ES complex (bottom) of K. oxytoca NiARD. Inserts show an expansion of the spectrum around 8332 eV where the 1s → 3d transition is expected. Bottom: Ni K-edge Fourier-transformed (k 2 – 12.5 Å1) EXAFS spectra (left), and Fourier-filtered (back-transform window 1.1 – 4.2 Å) EXAFS spectra (right) of the resting (top), and the ES complex (bottom) of K. pneumoniae NiARD. Data points are represented by open circles and the fit by a solid line. Reproduced by permission in part from [7]. Copyright (2002) American Chemical Society.

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Ni-ARD + substrate

Ni-ARD

80.0

70.0

60.0 ppm

50.0

40.0

Figure 6. Hyperfine-shifted resonances in the 14 T (600 MHz) 1H NMR spectra of 1 mM Ni-ARD in 90% H2O/10% D2O (pH 7.4 d-tris 10 mM) without substrate (bottom), and with 10 mM substrate added (top). All spectra were obtained at 298 K.

sufficiently flexible region of the structure (H96-T97-H98-G99), and is the most likely candidate for displacement by substrate. Direct ligation of substrate by Ni2 is also supported by paramagnetic NMR studies. A series of hyperfine-shifted resonances in the downfield region of the 1H NMR spectrum of resting state NiARD have been assigned to histidine imidazole protons further than 3 bonds from the nickel ion (Figure 6), and the pattern is consistent with three His ligands for the Ni2, two ligating through the ε2 N and one through the δ1 N in the resting state enzyme [7]. Addition of acireductone to the sample results in splitting of these peaks into a more complex pattern, clearly indicating a direct perturbation of the metal center by substrate [7]. A model for the NiARD active site, the hydroxamate complex 10 (see Figure 8 below), has been synthesized and characterized, and exhibits NMR hyperfine shift patterns similar to that found in NiARD [46]. Non-averaged dipolar coupling between unpaired electron spins and nuclear spins results in efficient relaxation of the nuclear spins. In the case of NiARD, this results in very broad lines for protons within ⬃9 Å of the metal center, and renders standard 1H detected multinuclear NMR methods useless in the vicinity of the active site. However, lower gamma nuclei such as 15N and 13C can still be detected directly, due to their weaker dipolar interactions with unpaired electrons than 1H under similar conditions. Thus, direct 13C detection 2D NMR methods offer the promise of sequential resonance assignments and structural insights into the active sites of ARD and related proteins. We have described the use of 2D Met. Ions Life Sci. 2, 473–500 (2007)


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multiple-quantum spectroscopy to detect 13C(O)-15N and 13C(O)-Cα correlations in the NiARD active site [47], and other researchers have published experiments that offer the hope of direct structural methods for determining structures of metal binding sites in paramagnetic proteins [48–51].

7.

ENZYMATIC STUDIES OF ACIREDUCTONE DIOXYGENASE

To date, the only comprehensive examination of mechanistic enzymology for ARD is the paper by Dai et al. [5]. A detailed discussion of the aqueous chemistry of acireductone substrates for ARD is provided by Zhang et al. [28], and that study has insights useful for considering the mechanism of ARD enzymes. Both the Ni and Fe forms of K. oxytoca ARD are reasonably efficient enzymes (kcat 5.0 102 s1 and 2.6 102 s1, respectively [5]). Kinetic analyses indicate a sequential mechanism is operating for both forms of the enzyme, and both substrates, acireductone and dioxygen, must bind to the enzyme prior to substrate release. Furthermore, substrate binding is ordered; in the absence of acireductone, O2 does not bind to either form of the enzyme in significant quantities, effectively ruling out a direct O2-M2 bound intermediate. However, acireductone binds stoichiometrically to the enzyme under anaerobic conditions, and binding is accompanied by a red shift of the acireductone UV–visible spectrum from 305 to 345 nm, indicating that the acireductone binds to the Ni2 as a dianion [5]. The strictly conserved Arg 104 is appropriately positioned to interact with substrate bound to the Ni2, and may also serve as a proton acceptor for the deprotonation that accompanies substrate binding (see Figure 4d). As might be expected based on the exposed nature of the active site, ARD is rather tolerant of different substituents at the acireductone functionality: modified acireductones with phenyl, propyl and methyl substituents at C-3 of the acireductone are all substrates for ARD, although the phenyl- and methyl-substituted substrates are turned over more slowly than the propyl form. Some mechanistic insight is gained from the use of a cyclopropyl-substituted acireductone (7b). The cyclopropyl ring would be expected to open in the presence of a radical on C-3 if such a species is formed during the reaction cycle. This would give rise to a reactive radical species on the terminal C-6 of the ringopened species, which could presumably react with amino acid residues within the active site, resulting in suicide inactivation. In the absence of O2, the binary ES complex between ARD and substrate 7b is stable, indicating that a radical is not present. However, once dioxygen is introduced, the enzyme slowly undergoes irreversible inactivation in the presence of substrate 7b (⬃100 turnovers for ARD, and 20 turnovers for ARD′). This indicates that either a radical is involved in the reaction mechanism or that a radical side-reaction takes place occasionally [5]. Met. Ions Life Sci. 2, 473–500 (2007)

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MECHANISTIC CONSIDERATIONS: WHAT IS THE ROLE OF NI(II) IN ACIREDUCTONE DIOXYGENASE ACTIVITY?

Octahedral Ni2 has a reduction potential of 1.16 V compared with SHE in complexes involving N/O ligands [52]. As such, it is not surprising that ARD is the only known mononuclear Ni-based dioxygenase. The nickel bound in the ARD active site does not ligate O2 in the absence of substrate [5], and the acireductone substrate is sufficiently reactive towards dioxygen that O2 activation is probably unnecessary. Nevertheless, a bound metal is required in the ARD active site for catalytic activity, and Ni2(or Co2) binding is necessary and sufficient to give rise to the off-pathway chemistry leading to carbon monoxide formation from substrate in WT KoARD as well as a number of active site mutants of this enzyme [34]. At least in NiARD, the substrate is directly ligated to the metal in the ES complex, based on NMR and XAS data [7]. The wide range of redox potentials for the metals that give rise to ARD activity makes it unlikely that the metal undergoes even transient redox chemistry during the reaction(s). Both ARD and ARD′ are EPR-silent, and no evidence for longlived organic radicals have been observed, although as noted above, the radicalbased suicide inhibitor substrate 7c slowly inactivates both forms of the enzyme [5]. We have proposed that the metal acts as a regioselective activator for the substrate via the appropriate Lewis acid interactions with substrate oxygens [5,8]. The proposed mechanisms are shown in Figure 7. For both enzymes, the first step is deprotonation of substrate, leading to the formation of the dianion, followed by single electron transfer from substrate to O2, giving rise to a superoxide/ organic radical pair. The radical pair collapses by attack of superoxide at C-1 of the substrate. At this point, the reaction pathways diverge, depending upon the nature of the coordination of substrate to the metal. For the Ni-bound enzyme, the substrate is proposed to bind to the Ni2 as a bidenate ligand via the oxygens at C-1 and C-3, while for the Fe-bound form, complexation is via the oxygens at C-1 and C-2 [8]. In both cases, the ligation is cis in the metal coordination sphere. The nature of the ligation directs the formation of the cyclic peroxide species by attack at the electrophilically activated carbon, C-3 in the case of NiARD, leading to the formation of the five-membered ring intermediate. In the case of FeARD, C-2 is activated and a four-membered ring is formed. Both species then undergo concerted electrocyclic ring cleavage to yield either the off-pathway products (including CO) or the on-pathway product 8 (Figure 1) and formate. A functional model for the NiARD reaction that supports most of the features of the proposed mechanism has recently been described by Berreau and coworkers (Figure 8) [53]. The sterically bulky acireductone 11 was used to prepare an octahedral Ni(II) complex 12. The complex was crystallized under anaerobic conditions, and shows the predicted 1,3 cis-ligation of the acireductone to the nickel ion, although the substrate is only monoanionic rather than dianionic. Anaerobic addition of tetramethylammonium hydroxide resulted in the formation Met. Ions Life Sci. 2, 473–500 (2007)


O

O2

1e–

OH

O2–

O

O– O

M

–O O

R

O–

R

491

O O–

R

M

O

M

A -O

NiARD

O rotation

O O δ+

R

FeARD′

–O

O

O

–O

–

O–

R

Ni 2+

Ni 2+

O

O

O

2e– δ+

R

O– Fe 2+

O

–O

O

O

O O

R

O O

O–

O–

R O

O–

B

C O

RCOO– +

CO

ARD products

+

HCOO–

R

COO–

+

HCOO–

ARD′ products

Figure 7. Proposed mechanism for differential product formation by NiARD and FeARD. The first step in both cases is electron transfer from the doubly deprotonated acireductone to O2, resulting in the formation of superoxide. Nucleophilic attack at C-1 of the acireductone forms the common intermediate A. In the case of the nickel-containing enzyme, the metal is proposed to bind A via ligation at the 1 and 3 position oxygens, while the Fe-containing form ligates A via the 1 and 2 positions. In either case, Lewis acid activation by the metal to the carbon bonded to the ligating oxygen results in nucleophilic attack at the appropriate carbon (C-3 in NiARD, C-2 in FeARD) giving rise to either the five-membered intermediate B (NiARD) or the four-membered C (FeARD). In both cases, electrocyclic rearrangement as indicated by arrows then yields products. Bold face oxygens represent the results of isotopic labeling experiments to confirm the disposition of oxygen atoms from dioxygen in the product mixtures [5].

of a second as yet uncharacterized species accompanied by a UV–visible red shift similar to what is seen in the dianionic form of acireductone bound to enzyme [5]. This species reacts rapidly with molecular oxygen to yield CO as well as a product complex 13 containing two benzoate ligands resulting from the acireductone Met. Ions Life Sci. 2, 473–500 (2007)

492

OH O

N O

ClO4–

N Ni

N

O

OH

N N

11

10

HO Ph

Ph O

O2, Me2NOH

ClO4–

N

O Ni

CH3CN

N

N N

12 H N

N O

Ph Ph

O

H N O

Ni

O

O

Ph

+

C

O

Ph

N

13

Figure 8. Structural model 10 for the NiARD active site from [46]. See text for further details. Structure 12 is a mechanistic model for the active site of NiARD prepared using bulky acireductone 11. Upon treatment of 12 with base and oxygen, carbon monoxide is formed, along with complex 13. Both structures 12 and 13 have been characterized crystallographically [53]. Met. Ions Life Sci. 2, 473–500 (2007)


493

cleavage. Notably, this complex contains one fewer N-ligand than complex 12, which supports the proposed histidine ligand displacement proposed previously for KoARD [7]. Recently, another instance of Ni2 ligated by three histidine residues supporting oxidative chemistry has been reported in copper amine oxidase (CAO) [54]. The CAO pro-enzyme normally self-processes to form the co-factor 2,4,5-trihydroxyphenylalanine quinone from a Tyr residue in the active site. It was found that Ni2 supported the formation of the co-factor in the presence of molecular oxygen, but again, no redox state changes of the nickel was detected, and direct electron transfer between the Ni(II)-tyrosinate species from molecular oxygen was proposed as the rate-limiting step in the reaction [54].

9.

STRUCTURALLY AND FUNCTIONALLY RELATED ENZYMES

The ligand geometry found in the ARD active site has proven to be a common motif used by cupins for binding a variety of divalent metal ions [41,42,55,56]. The HXHXXX(X)E motif that provides three of the four protein-based ligands in ARD is conserved in many cupin metalloproteins, with the third His ligand occurring typically approximately 30–40 residues towards the C-terminal of the polypeptide (see Figure 9). This motif appears to provide both chelative stability for tight metal binding combined with sufficient flexibility to allow changes in ligation geometry and dentation (one or two ligands) for metal-substrate interactions. This flexibility might be provided by either slight changes in ligand positions or by a change in which imidazole nitrogen (Nε1 vs Nδ2) is used for metal ligation, although no direct structural evidence of the imidazole ligand switch has been found to date. Cupins using this metal binding motif include oxalate oxidase (germin) [56] and oxalate decarboxylase [57], both of which coordinate Mn2 in an octahedral ligation sphere with two open cis valence sites occupied by non-protein ligands. Both of these enzymes catalyze reactions of oxalic acid. Oxalate oxidase uses one O2 and one oxalate to produce two molecules of CO2 and one of hydrogen peroxide. Oxalate decarboxylase catalyzes the disproportionation of oxalate without O2, producing a molecule of formate and a molecule of CO2. Even more closely related to ARD in terms of the overall chemistry that is catalyzed are the quercetin 2,3-dioxygenases (QDs). These enzymes are also members of the cupin superfamily, and catalyze the reaction shown in Figure 10. Two forms of QDs have been identified, a Cu2-dependent form and a Fe2dependent form. In the anaerobic structure of the copper-containing QD from Aspergillus japonicus, the copper ion is coordinated by three His and one Glu, but only one valence site is available to bind substrate, resulting in a penta-coordinate copper ligation that is square pyramidal distorted toward trigonal bipyramidal Met. Ions Life Sci. 2, 473–500 (2007)

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Figure 9. Active sites of metalloenzymes from the cupin superfamily. Top row: NiARD from K. oxytoca (PDB entry 1ZRR [43]) MmARD from M. musculus (PDB entry 1VR3 [57]), with unidentified metal. Second row: differential iron binding in quercetin dioxygenase from B. subtilis (PDB entry 1Y3T [60]). Third row: copper-containing quercetin dioxygenase from A. japonicus (PDB entry 1H1I [59]). Fourth row: Mn2-containing cupins oxalate oxidase (germin), (PDB entry 1F12 [56]) and oxalate decarboxylase, PDB entry 1H58 [57]). Met. Ions Life Sci. 2, 473–500 (2007)


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OH

OH OH

OH

O

HO

O QD

O2

* OH

HO

CO O

OH

COOH

O OH

Figure 10. O2 oxidation of quercetin catalyzed by quercetin 2,3-dioxygenase. Starred (*) carbon is released as CO.

[58]. In the absence of substrate, the Cu2 appears to be mixed between a fourand five-coordinate state with water as one ligand and the carboxylate ligand partially dissociating from the copper ion [59]. The quercetin substrate appears to be distorted upon binding to the enzyme, leaving open the possibility that the spin-forbidden nature of the reaction between dioxygen and quercetin may be relieved by an induced partial radical character in the bound substrate, resulting either from spin delocalization onto the Cu2 or the structural distortion (or both). A similar situation is found for the active sites of a Fe2-containing QD from B. subtilis, a functional dimer [60]. In this case, while the Fe2 ions are pentacoordinate in both active sites, one active site contains the glutamate carboxylate in close coordination with the iron, while in the second site, the glutamate-Fe distance is greater, and the trigonal bipyramidal arrangement of ligands is slightly distorted. While direct interaction between metal and O2 has been proposed in the case of the copper-containing QD [58], the use of NO as an oxygen mimic in EPR studies of the Fe-containing form suggests that O2 would be shielded from the Fe2 center in the substrate-bound form of QD [60]. To date, therefore, no direct evidence for O2 activation by metal ligation has been found for either QD or ARD.

10.

FUTURE DIRECTIONS

The most pressing unanswered questions relating to ARD can be summarized briefly as follows. (1) What is the structural switch responsible for the change in activity that accompanies the change in metal between ARD and ARD′? (2) Do vertebrates (and in particular, humans) incorporate the off-pathway chemistry catalyzed by NiARD under normal or abnormal physiological conditions? (3) What is the native metal found in human ARD as isolated from tissue? Dealing with these questions in the order that they are presented, we first consider the structural switch accompanying the change in metal at the active site of Met. Ions Life Sci. 2, 473–500 (2007)

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ARD. As we have already noted, replacing Ni2 with Fe2 or vice versa is necessary and sufficient to obtain the enzymatic activity associated with that metal. The chromatographic separability of the two enzymes implies a significant structural difference between the two forms, and indeed, comparison of 1H,15N HSQC spectra of the two forms of the enzyme suggest substantial structural differences between them [34]. We are currently in the process of refining a structure for an FeARD structural homolog, and have found that the differences are indeed extensive, and are to be found primarily in regions that do not form the cupin barrel. This work will be published shortly. Still, the question of whether the metal itself drives the different chemistries or whether changes in ligand or second-sphere interactions are responsible for the divergent chemistry remains an open question. Mutagenesis results thus far have been inconclusive in this regard, but mutation of the most likely ligand switch, the relatively mobile His 98, interferes with binding of both Fe2 and Ni2, and is thus unlikely to be the deciding factor in the structural switch. The possibility for off-pathway activity by a human ARD homolog, although so far unsuccessful in vivo, is particularly interesting for a number of reasons. First, the off-pathway reaction product, carbon monoxide (CO), has been implicated as a signaling molecule in a variety of cellular processes. A role for CO similar to that of NO in the activation of adenylate cyclase was suggested several years ago [61]. The role of CO as an anti-apoptotic has been shown in a variety of clinical and pharmacological settings [62–64], and there is evidence for stimulation of angiogenesis by carbon monoxide produced endogenously [65]. A SAGE analysis [66] of mRNA levels corresponding to the human homolog of ARD, SipL, shows the highest levels of expression in tumor cell lines associated with breast cancer tissues, with lesser levels in normal liver and stem cell tissues. However, the SAGE assay does not differentiate between the N-truncated SipL and the full-length MTCBP-1, and expression of the full-length MTCBP-1 appears to be suppressed in other tumor lines, perhaps due to the regulatory role of this protein in controlling the activity of matrix metalloproteinases [38]. We have found that the bacterially expressed human HsARD homolog (MTCBP1) does indeed function as an ARD in vitro, and that both on-pathway and offpathway activity is observed from different fractions of the bacterially expressed HsARD (G. Pagani, unpublished results). The discovery that MTCPBP-1 can act to restore viability in yeast knockout strains strongly supports a role for MTCPBP1 in the methionine salvage pathway. However, recovery of sufficient amounts of the enzyme from mammalian tissues for metal analysis has not yet been possible, and the nature of the metal bound in vivo has not yet been established. Still, the observation of both types of activity in vitro leaves open the possibility that the off-pathway ARD chemistry has physiological relevance in humans, either under normal conditions or in pathology, and that a human Ni-containing enzyme may yet be identified in vivo.

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ACKNOWLEDGMENTS T.C.P. acknowledges support of the US Public Health Service (R01-GM-067786). G.M.P. acknowledges previous support by USPHS training grant GM007596.

ABBREVIATIONS Ado-Met ALP ARD CAO EDTA ES EXAFS HsARD HSQC KoARD MmARD MSP MT1-MMP MTA MTAN MTAP MTCBP-1 MTR-1-PI MTRK ODC ORF OsARD Pi QDs RCSB RuBisCO SAGE SAM Sip2 SipL XANES XAS

S-adenosylmethionine ( SAM) acireductone dioxygenase-like protein acireductone dioxygenase ( 2-hydroxy-3-keto-5-thiomethylpent1-ene dioxygenase) copper amine oxidase ethylenediamine-N,N,N′,N′-tetraacetate enzyme-substrate extended X-ray absorption fine structure spectroscopy Homo sapiens acireductone dioxygenase heteronuclear single quantum correlation Klebsiella oxytoca acireductone dioxygenase Mus musculus acireductone dioxygenase methionine salvage pathway membrane-type 1 matrix metalloproteinase methylthioadenosine methylthioadenosine nucleosidase methylthioadenosine phosphorylase membrane-type 1 matrix metalloproteinase cytoplasmic tail-binding protein-1 5-methylthioribose-1-phosphate isomerase 5-methylthioribose kinase ornithine decarboxylase open reading frame Oryza sativa acireductone dioxygenase ( Sip2) inorganic phosphate quercetin dioxygenases Research Collaboratory for Structural Bioinformatics ribulose-1,5-bisphosphate carboxylase/oxygenase serial analysis of gene expression S-adenosylmethionine ( Ado-Met) submergence-induced protein 2 ( OsARD1) submergence-induced protein-like protein X-ray absorption near-edge spectroscopy X-ray absorption spectroscopy

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13 The Nickel-Regulated Peptidyl Prolyl cis/trans Isomerase SlyD Frank Erdmann and Gunter Fischer Max-Planck Research Unit for Enzymology of Protein Folding, Weinbergweg 22, D-06120 Halle, Germany <[email protected]>

1. INTRODUCTION 2. SlyD BELONGS TO THE PEPTIDYL PROLYL cis/trans ISOMERASES 2.1. Reactions Catalyzed by SlyD 2.2. Repertoire of Peptidyl Prolyl cis/trans Isomerases in Escherichia coli (E. coli) 2.3. Distribution of slyD Genes across Organisms 2.4. Biochemical Information about E. coli SlyD 2.4.1. Structural Information about SlyD 2.4.2. SlyD Exhibits Peptidyl Prolyl cis/trans Isomerase and Chaperone-like Activity 2.4.3. Ni2⫹ Ions Inhibit the Peptidyl Prolyl cis/trans Isomerase Activity of SlyD 2.4.4. SlyD Binds Nucleotides in a Metal Ion-dependent Manner 2.4.5. SlyD Inhibits Mammalian Adenylate Cyclase 3. INSIGHTS INTO THE BIOLOGICAL ROLE OF SlyD 3.1. Lysis of E. coli by Bacteriophage ΦX174 Requires slyD 3.2. SlyD is Linked to the Hydrogenase Biosynthetic Pathway



502 503 503 504 505 508 508 511 511 512 513 513 513 514

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4.

CONCLUSIONS ABBREVIATIONS AND DEFINITIONS REFERENCES

1.

INTRODUCTION

515 515 516

E. coli SlyD (also known as E. coli FKBP21 or WHP [1]) and its homologs are Ni2⫹-binding enzymes encompassing two distinct domains. The domains consist of a peptidyl prolyl cis/trans isomerase (PPIase) consensus sequence element and a Cterminal histidine/cysteine-rich extension (CTE) capable of binding bivalent metal ions [2]. Acceleration of prolyl bond cis/trans isomerization by SlyD in the absence of Ni2⫹ ions revealed the formation of a functional PPIase site and accessibility of the active site to oligopeptide and protein substrates. The enzyme can monitor the presence of Ni2⫹ ions by becoming reversibly inhibited. At present, SlyD represents the only known example of a transition metal ion-regulated PPIase. The Zn2⫹dependent folding-arrest activity of the Thermus thermophilus trigger factor, which represents a ribosome-bound FKBP, appears to be independent of its PPIase activity [3]. Although many PPIases are known to be present in E. coli, enzymes of the SlyD type might contribute to the control of the slow rotational movement underlying prolyl bond cis/trans isomerization in specific substrates (Figure 1). Generally, the free energy dependence of the reaction progress of a peptide bond cis/trans isomerization in a polypeptide chain segment indicates just two minimum-energy structures. Nineteen out of the twenty proteinogenic amino acids form secondary amide peptide bonds. In contrast, an imidic peptide bond is created exclusively by the N-alkylated amino acid proline on its N-terminal acylation site. Due to their unique imidic structure, prolyl bonds have a particular electron distribution, and cis and trans prolyl bond isomers suffer from similar

O

O

C N

Figure 1.

O C ω

SlyD

The two isomeric states of a prolyl bond.

Met. Ions Life Sci. 2, 501–518 (2007)

C N

C ω

O

Ni-REGULATED PEPTIDYL-PROLYL CIS/TRANS ISOMERASE

503

steric constraints that lead to comparable free energy for both conformational states. The conformational isomers at the angles ω ⯝ 0⬚ (cis) and ω ⯝ 180⬚ (trans) are separated by a rotational barrier ∆G‡ of about 80 kJ/mol, corresponding to the energy change of the perpendicular arrangement of the carbonyl plane to the prolyl ring (ω ⯝ 90⬚) relative to the planar ground states [4]. Remarkably, this conformational interconversion is often found to be the ratelimiting step in protein folding, and has been shown to be important for the conformational control of the biological activity of proteins [5,6]. Furthermore, the results of kinetic experiments frequently point to a very high rotational barrier between both conformational states of peptidyl prolyl bonds in all folding states of a polypeptide backbone. Under physiological conditions this barrier cannot be easily diminished by environmental parameters or small molecule catalysts. The electronic force by which C–N bonds may confer rotational uncoupling of elementary processes during folding is through peptide bond rigidity bordered by freely rotatable single bonds. Thus, the properties of this bond have an important influence on backbone stiffness at an intramolecular level in all folding states of a protein and determine the thermodynamics of intermolecular protein–protein and protein–ligand interactions.

2. SlyD BELONGS TO THE PEPTIDYL PROLYL cis/trans ISOMERASES 2.1.

Reactions Catalyzed by SlyD

The enzyme repertoire of all living organisms has been found to serve as a source of specific and efficient catalysts for prolyl isomerization in various substrates, and SlyD is among them. The first report of PPIase catalysis indicated the existence of these enzymes in the porcine kidney cortex [7], and the enzyme class (EC 5.2.1.8) was named according to their catalyzed reactions—the cis to trans and trans to cis isomerization of prolyl bonds. Peptide bond cis/trans isomerases represent the only example of biocatalysts directed toward a conformational interconversion in a protein known to date. PPIases evolved early during the course of evolution, and their three different enzyme families probably derived from a common ancestor. Currently, PPIases are subdivided into the subfamilies of cyclophilins (Cyp), FK506-binding proteins (FKBP), and parvulins (Par) (Figure 2). The family members are unrelated to each other in their amino acid sequences, have distinct substrate specificities, and are sensitive to different types of inhibitors. All enzymes of the SlyD type belong to the FKBP family of PPIases. FKBPs are ubiquitous and abundant enzymes which have been detected in higher and lower eukaryotes as well as in prokaryotic organisms. In most organisms there is a multiplicity of FKBPs, differing in domain composition, localization, substrate Met. Ions Life Sci. 2, 501–518 (2007)

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specificity, and cellular expression level. Functional overlap within as well as between the families is likely to occur in vivo. Specific members of the cyclophilin, FKBP, and parvulin families have been shown to play a pivotal role in intracellular signalling, bacterial virulence, cell cycle integrity, viral infections, receptor recognition, and channel modulation. Biochemically, prototypical PPIases exert their function on the basis of several reaction types which might control, either in isolation or collectively, the physiological processes driven by these enzymes [5]. Among them, the catalysis of prolyl isomerization and a holding function for unfolded proteins seem to be important for SlyD. The enzymatic activity of PPIases is routinely assessed by an assay based on accelerated reaction rates in the presence of PPIase for the isomer-specific proteolysis of tetrapeptide substrates with the common sequence Suc-Ala-Xaa-Pro-Yaa4-nitroanilide [8,9]. Another standard assay is based on the acceleration of the slow kinetic phases during the refolding of denatured RNase T1 [10].

2.2.

Repertoire of Peptidyl Prolyl cis/trans Isomerases in Escherichia coli

Since most investigations of SlyD have been carried out with the E. coli enzyme, the whole spectrum of PPIases in this organism should be considered (Figure 2).

Figure 2. Repertoire of PPIases in E. coli. The enzyme class of PPIases is subdivided into three independent families, the cyclophilins, the FKBPs and the parvulins. The name of the respective PPIase gene is given in italics.

Met. Ions Life Sci. 2, 501–518 (2007)


505

Most of these proteins are catalytically efficient enzymes in a standard PPIase assay. Only two cyclophilins were found in E. coli, the cytoplasmic EcCyp18 (PpiB) and the periplasmically localized EcCyp20 (PpiA) [11,12]. The catalytic activity of these prototypic cyclophilins is superior to mammalian Cyp18. In contrast, cyclosporin A, which is inhibitory in the low nanomolar range toward human Cyp18, has only low affinity for E. coli cyclophilins [13]. FKBPs constitute the largest family of E. coli PPIases, in that five members have been identified in the E. coli genome. This family includes the small EcFKBP16 (also termed SlpA), SlyD (rationally termed EcFKBP21), which are both Mip-related enzymes, the cytoplasmic EcFKBP22 (FklB) and periplasmic EcFKBP29 (FkbA), and the ribosome-associated EcFKBP48, commonly termed the ‘trigger factor’ [14]. The prototypic parvulin, the EcPar10, was purified and characterized by Rahfeld et al. [15]. Larger parvulins, the SurA and the PpiD, seem to be multifunctional enzymes [16–18]. Genetic inactivation of the four periplasmic PPIases of E. coli, the two parvulin-like PPIases, the Mip-like FKBP and a cyclophilin, resulted in a strain viable under laboratory conditions. However, severely defective pilus production might reduce survival rates under environmental stress [19].

2.3.

Distribution of slyD Genes across Organisms

SlyDs are ubiquitous among prokaryotes, in that genes exist that encode proteins with a putative metal ion-binding domain attached C-terminal to a FKBP domain. Typically, a low level of amino acid sequence similarity exists between prototypic human FKBP12 and the FKBP-domain of SlyD proteins. For E. coli SlyD a sequence identity of 28.1% was calculated (Table 1). Fifty-nine proteins from 47 different species of prokaryotes revealed amino acid sequence similarity to E. coli SlyD. Apparently, the maximum number of slyD-like genes placed in an individual genome was two. Moreover, SlyDs of different species share a similar domain arrangement (Figure 3). Eukaryotes seem to lack a PPIase capable of attracting metal ions for regulation of enzyme activity. As an exception, the enzyme function of human FKBP38 is indirectly controlled by Ca2⫹ ions using calmodulin as mediator molecule [20]. Originally, authentic SlyD was isolated from E. coli lysates. Over the last few years, many SlyD homologs have been identified by genome or proteome analyses (Table 1). SlyD sequences encode proteins with a molecular mass in the range 17–35 kDa. It is noteworthy that the SlyD from Haemophilus influenzae (H. influenzae) appears to be the only FKBP gene in the genome of that organism [21]. The conservation of slyD in the small H. influenzae genome suggests that SlyD may be an important component in FKBP-dependent protein maturation machinery.

Met. Ions Life Sci. 2, 501–518 (2007)

Swiss-Prot/TrEMBL entry Q6FCZ8_ACIAD O28290_ARCFU Q5NYP1_AZOSE Q6MPW7_BDEBA Q6MQW6_BDEBA Q4HFP0_CAMCO Q9PJ12_CAMJE Q5HX51_CAMJR Q4HM23_CAMLA Q4HQY7_CAMUP Q7NS45_CHRVO Q6CZU7_ERWCT SLYD_ECOLI (P0A9K9) SLYD_ECO57 (P0A9L1) SLYD_ECOL6 (P0A9L0) Q7VP93_HAEDU SLYD_HAEIN (P44830) Q4QMM1_HAEI8 Q5V2S8_HALMA Q9P9H4_HALSA Q7VJ14_HELHP SLYD_HELPY (O25748) SLYD_HELPJ (Q9ZK89) Q5R0Z5_IDILO Q72RL4_LEPIC Q6LZQ2_METMP Q8TM55_METAC Q60CM5_METCA Q7DDB0_NEIMB Q82U07_NITEU Q9CL91_PASMU Q7N9C1_PHOLL

Organism

Acinetobacter specialis (strain ADP1) Archaeoglobus fulgidus Azoarcus specialis (strain EbN1) Bdellovibrio bacteriovorus Bdellovibrio bacteriovorus Campylobacter coli (strain RM2228) Campylobacter jejuni Campylobacter jejuni (strain RM1221) Campylobacter lari (strain RM2100) Campylobacter upsaliensis (strain RM3195) Chromobacterium violaceum Erwinia carotovora subsp. atroseptica* Escherichia coli Escherichia coli (strain O157:H7) Escherichia coli (strain O6) Haemophilus ducreyi Haemophilus influenzae Haemophilus influenzae (strain 86-028NP) Haloarcula marismortui* Halobacterium salinarium* Helicobacter hepaticus Helicobacter pylori* Helicobacter pylori (strain J99)* Idiomarina loihiensis Leptospira interrogans* Methanococcus maripaludis Methanosarcina acetivorans Methylococcus capsulatus Neisseria meningitidis (serogroup B) Nitrosomonas europaea Pasteurella multocida Photorhabdus luminescens subsp. laumondii

ACIAD1183 AF1989 AZOSEA36980 Bd0722 Bd0336 CCO1766 Cj0115 CJE0110 CLA0559 CUP1387 CV3582 ECA4054 b3349 z4707 c4123 HD0202 HI0699 NTHI0822 rrnAC1230 VNG1294G HH0437 HP1123 JHP1052 IL0821 LIC11731 MMP0572 MA2813 MCA0081 NMB1522 NE1706 PM1349 plu0422

Ordered locus/ORF names 17152 Da 28952 Da 17575 Da 17765 Da 17177 Da 20091 Da 20132 Da 20160 Da 20558 Da 19551 Da 17415 Da 20820 Da 20853 Da 20853 Da 20853 Da 20769 Da 20658 Da 20644 Da 34839 Da 33345 Da 19015 Da 19997 Da 20100 Da 17499 Da 17853 Da 25537 Da 28176 Da 17316 Da 17405 Da 17841 Da 25798 Da 20203 Da

Molecular weight

Table 1. Distribution of slyD across organisms. Data collected from the Swiss-Prot Database (release 48.1 of 27 September 2005) and TrEMBL Database (release 31.1 of 27 September 2005).

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Met. Ions Life Sci. 2, 501–518 (2007)

Q7MV03_PORGI Q9I5A3_PSEAE Q88PS2_PSEPK Q4FV05_PSYAR Q9V0N6_PYRAB Q8Y095_RALSO Q57J17_SALCH Q5PL23_SALPA Q8Z1Y6_SALTI Q8ZLL4_SALTY Q8EBT2_SHEON SLYD_SHIFL (P0A9L2) Q3YWS3_SHISO SLYD_TREPA (O83369) Q480G8_COLP3 Q47WN4_COLP3 Q7MA15_WOLSU Q8PIZ2_XANAC Q8PMR8_XANAC Q8P7M9_XANCP Q8PB16_XANCP Q5GXM7_XANOR Q5H1M4_XANOR Q87BD7_XYLFT Q74XZ9_YERPE Q8ZJC2_YERPE

*Erwinia carotovora subsp. atroseptica ⫽ Pectobacterium atrosepticum Haloarcula marismortui ⫽ Halobacterium marismortui Halobacterium salinarium ⫽ Halobacterium halobium Helicobacter pylori ⫽ Campylobacter pylori Leptospira interrogans ⫽ Leptospira interrogans serogroup Icterohaemorrhagiae serovar copenhageni Porphyromonas gingivalis ⫽ Bacteroides gingivalis Ralstonia solanacearum ⫽ Pseudomonas solanacearum Vibrio psychroerythus ⫽ Colwellia psychrerythraea (strain 34H / ATCC BAA-681)

Porphyromonas gingivalis* Pseudomonas aeruginosa Pseudomonas putida (strain KT2440) Psychrobacter arcticum Pyrococcus abyssi Ralstonia solanacearum* Salmonella choleraesuis Salmonella paratyphi A Salmonella typhi Salmonella typhimurium Shewanella oneidensis Shigella flexneri Shigella sonnei (strain Ss046) Treponema pallidum Vibrio psychroerythus* Vibrio psychroerythus* Wolinella succinogenes Xanthomonas axonopodis pv. citri Xanthomonas axonopodis pv. citri Xanthomonas campestris pv. campestris Xanthomonas campestris pv. campestris Xanthomonas oryzae pv. oryzae Xanthomonas oryzae pv. oryzae Xylella fastidiosa (strain Temecula 1) Yersinia pestis Yersinia pestis PG1315 PA0837 PP0776 Psyc_0283 PAB1864 RS04602 SC3389 SPA3321 STY4343, t4050 STM3455 SO3417 SF3367 SSO_3479 TP0349 slyD1: CPS_2846 slyD2: CPS_4133 WS0525 no data available no data available no data available no data available XOO3290 XOO1893 PD1519 YP0191 YPO0193, y3975

20980 Da 17010 Da 17206 Da 17426 Da 30019 Da 19405 Da 20788 Da 20788 Da 20848 Da 20788 Da 23491 Da 20853 Da 20853 Da 18428 Da 18359 Da 17610 Da 18836 Da 17444 Da 16751 Da 17497 Da 16699 Da 17456 Da 18627 Da 17514 Da 22011 Da 21052 Da

507

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ERDMANN and FISCHER PPIase domain

MW

CTE

20.8 kDa I: 87 %

S: 94 %

I: 67 %

S: 73 %

20.8 kDa I: 68 %

S: 79 %

I: 54 %

S: 61 %

20.7 kDa I: 38 %

S: 53 %

I: 22 %

S: 27 %

17.8 kDa I: 33 %

S: 47 %

I: 26 %

S: 31 %

17.4 kDa I: 30 %

S: 47 %

I: 24 %

S: 27 %

17.8 kDa I: 48 %

S: 59 %

I: 22 %

S: 26 %

17.2 kDa I: 97 %

S: 99 %

I: 87 %

S: 89 %

20.8 kDa I: 100 % S: 100 %

I: 100 % S: 100 %

20.8 kDa I: 42 %

S: 51 %

I: 20 %

S: 24 %

17.5 kDa I: 41 %

S: 51 %

I: 22 %

S: 24 %

17.4 kDa I: 76 %

S: 81 %

I: 67 %

S: 71 %

22.0 kDa

Figure 3. Domain composition of SlyD proteins from selected organisms. In comparison with E. coli SlyD amino acid sequence identity (I) and similarity (S) are given for the FKBP-like domain (light grey) and the CTE domain (dark grey).

2.4. Biochemical Information about E. coli SlyD 2.4.1.

Structural Information about SlyD

E. coli SlyD was first characterized using two different approaches in 1994 [1,22], and finally named according to its function to mediate phage protein E-directed Met. Ions Life Sci. 2, 501–518 (2007)


509

flap

? bulge CTE

FKBP12

SlyD

Figure 4. The three-dimensional structure of human FKBP12 (PDB: 1FKF), MtFKBP17 (PDB: 1IX5), and the putative structure of SlyD as predicted by SWISS-MODEL [45]. The SlyD contains a probably disordered CTE.

lysis of bacterial cells (sensitive to lysis D). It consists of two domains, a Nterminal FKBP domain of about 150 amino acids and a C-terminal metal-binding motif of about 50 amino acids (Fig 3). Independent of metal ions, SlyD has a tendency to dimerize in solution [23]. The three-dimensional structure of SlyD is currently unknown. However, structural data on numerous prokaryotic and eukaryotic FKBPs are available and allow the prediction of the secondary and tertiary structure of SlyD by computational analysis. When comparing the amino acid sequences of E. coli SlyD and FKBPs for which the dimensional structure is already known, the highest level of sequence similarity of 40% was found with the 17 kDa FKBP from Methanococcus thermolithotrophicus (MtFKBP17). Based on the NMR solution structure of MtFKBP17 [24], the tertiary structure of SlyD was modelled by SWISS-MODEL and is shown, together with the structures of MtFKBP17 and the prototypic hFKBP12, in Figure 4. A structure-based sequence alignment of SlyD with the MtFKBP17 and hFKBP12 was also performed to identify differences in the secondary structure of the FKBP (Figure 5). A dominant feature of both structures is the long insertion (IF) of 47 amino acid residues in the flap region between β2 and β3 strands that form a satellite-like subdomain tethered to the FKBP12-like module. This subdomain encompasses secondary structure elements with the topology of β6-α3β9-β8-β7, and exposes a hydrophobic surface that could establish a number of secondary binding sites for enzyme-substrate interactions. Apparently, the enlarged flap region is an essential structural component of this property. An IF segment was also identified in non-SlyD-like FKBP, such as in Methanococcus jannaschii, Pyrococcus horikoshii, and Aeropyrum pernix [24]. Met. Ions Life Sci. 2, 501–518 (2007)

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Figure 5. The structure-based sequence alignment of SlyD with human FKBP12 and MtFKBP17. Multiple alignments were carried out using CLUSTALW [46] and structurebased alignments were performed using DSViewer. Secondary structure elements unique to human FKBP12 are shown in black, while structure elements which can be found in all three proteins are white. Insertions in the flap and bulge regions, exclusively found in SlyD and MtFKBP17, are grey in color.

SlyD and MtFKBP17 lack the hFKBP12 typical N-terminal β1-strand. In the four-stranded β -sheet A and α1-helix containing PPIase domains of both proteins a topology of β4-β5-β1-β2-β3 was found, which agrees with the secondary structure of hFKBP12, except for the β1-strand. The β5-strand of SlyD and MtFKBP17 is divided into the β5a and β5b strands. In contrast to hFKBP12, the structure of SlyD was modelled to have a short α-helical insertion (α2) in the ‘bulge’ loop, similar to the MtFKBP17. The side chains of the amino acids of α2-helix form a hydrophobic patch together with the side chains on β4 and β5a-strand. Surprisingly, the β5a-strand of hFKBP12 is exclusively formed when an active site-directed ligand is bound. This implies that the α2-helices of SlyD might stabilize the β5a-strands, even in the absence of a ligand. The histidine/cysteinerich C-terminal extension of SlyD is devoid of hydrophobic amino acids and therefore presumably disordered. Met. Ions Life Sci. 2, 501–518 (2007)


2.4.2.

511

SlyD Exhibits Peptidyl Prolyl cis/trans Isomerase and Chaperone-like Activity

PPIase activity of SlyD has been determined under various conditions. Due to its high proteolytic sensitivity, SlyD did not show PPIase activity in the standard assay with α-chymotrypsin as auxiliary protease [1]. In contrast, SlyD remained sufficiently stable in the presence of trypsin or thrombin [2]. For this reason, PPIase activity measurements were carried out with trypsin as isomer-specific protease or in the protease-free PPIase assay according to Janowski et al. [25]. In the trypsin-coupled assay, which determines catalysis of the cis to trans isomerization, specificity constants (kcat /Km) of 29600 M⫺1s⫺1, 6200 M⫺1s⫺1, and 5600 M⫺1s⫺1 were determined for the substrates Suc-Ala-Phe-Pro-Arg-4-nitroanilide, SucAla-Ala-Pro-Arg-4-nitroanilide, and Suc-Ala-Leu-Pro-Arg-4-nitroanilide respectively. Using the protease-free PPIase assay, a Km value of 1.1 ± 0.6 mM and kcat ⫽ 16 ± 6 s⫺1 were calculated for the catalysis of cis to trans isomerization of Suc-Ala-Phe-Pro-Phe-4-nitroanilide. Due to a more favorable Km value, a protein substrate brings about almost a tenfold enhancement of catalytic power [26]. Catalytic activity of SlyD could not be inhibited by FK506 [2]. Two Michaelis complexes will exist as ground state polypeptide-binding structures on the catalytic pathway of SlyD and other PPIases. Complex formation indicates considerable affinity for unfolded or partially folded polypeptide chains of the enzymes [5]. In fact, Michaelis complex formation could involve prolineindependent secondary binding sites which might extend to distal portions of the active site [27]. Consequently, formation of the Michaelis complexes will allow the protection of client polypeptides from aggregation, thus assisting the proper folding of the clients. Apparently, this satellite segment between β2 and β3 strands builds a functional extension to the tethered catalytic site [24] that makes SlyD superior to FKBP12 in facilitating soluble expression of overproduced proteins. As an example, the membrane spanning subunit gp41 of the HIV-1 precursor protein gp160 could be expressed in a biologically active state in E. coli when covalently linked to a C-terminal truncated SlyD or tandem SlyD variant [28]. Notably, SlyD is a frequent contamination of proteins recombinantly expressed in E. coli, in particular those containing a Hisn tag, when purified by immobilized metal-ion affinity partitioning (IMAP) [23,29–31].

2.4.3.

Ni2⫹ Ions Inhibit the Peptidyl Prolyl cis/trans Isomerase Activity of SlyD

The C-terminal part of SlyD is rich in potentially metal-chelating amino acids, such as histidine, cysteine, glutamate, and aspartate, which enable the formation of protein–metal complexes with one or several metal ions. Furthermore, it is possible that different metal ions are bound to different residues according to their Met. Ions Life Sci. 2, 501–518 (2007)

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coordination requirements. The first detailed investigations of metal-binding properties were performed by Wülfing et al. [1]. In reconstitution experiments, they could show that SlyD has the highest affinity to Ni2⫹ and Zn2⫹ ions. Moreover, Cu2⫹ and Co2⫹ were also complexed, albeit to a lesser extent, probably due to their strict geometrical coordination requirements. In contrast, iron (neither as Fe2⫹ nor Fe3⫹) is not bound at all. The earth alkaline metals Mg2⫹ and Ca2⫹ cannot form complexes with SlyD either. In conclusion, the metal-binding domain of SlyD seems to be able to bind bivalent metal ions within a certain range of ionic radii. Hottenrott et al. [2] demonstrated that the PPIase activity of SlyD was decreased by 20 µ M Ni 2⫹ to a limiting value of 10% of the Ni 2⫹-free control. In contrast, millimolar concentrations of Na⫹, K⫹, Ca 2⫹, and Mg2⫹ ions did not affect the PPIase activity. The CTE domain conferred metal-ion sensitivity. A SlyD molecule contains three metal-binding sites at which Ni 2⫹ is fi rmly bound (K1, K2 , and K3 were calculated to be 9.5 ⫻ 10⫺5 M, 4.9 ⫻ 10⫺5 M, and 4.4 ⫻ 10⫺5 M, respectively). Nickel ions inhibited the PPIase activity of SlyD with a Ki value of 1.6 µ M. It seems likely that the inhibitory effect of Ni2⫹ parallels the binding of a single metal ion. A structural component of metal ion binding became obvious by far CD spectra of SlyD recorded in the presence and absence of Ni 2⫹. The results showed that [SlyD•3Ni2⫹] exhibits increased β -turn conformation in comparison to the nickel-free enzyme. These fi ndings do not answer the question to what extent SlyD exhibits PPIase activity in bacterial cells. Since there is a residual PPIase activity under Ni 2⫹-saturating conditions in vitro, changed substrate specificity for [SlyD•nNi 2⫹] can also be assumed.

2.4.4.

SlyD Binds Nucleotides in a Metal Ion-dependent Manner

A nucleotide-binding site could be identified in SlyD using 32P labeling experiments because various ATP analogs could be covalently linked to the protein [23]. A glycine-rich stretch, G181GEGCCGKG189, and a histidine triad, H149GHVHGA155, both typical of nucleotide-binding motifs, were hypothesized to constitute the nucleotide-binding site of E. coli SlyD. Covalent labeling of SlyD by ATP analogs was shown to depend on the presence of metal ions, indicating decreasing labeling yield in the series Zn2⫹ ⱖ Ni2⫹ ⬎ Co2⫹ ⬎ Cu2⫹, whereas Mg2⫹ and Ca2⫹ did not allow incorporation of 32 P. Furthermore, covalent labeling of SlyD by an ATP-derived photoaffinity ligand was suppressed by the addition of ATP, ADP, GTP, and UTP. However, the labeling does not reflect phosphorylation of the protein, because no radioactivity was incorporated when SlyD was incubated with [γ -32P]ATP. These results show that SlyD might be considered as a nucleotide-binding protein which interacts equally well with purine or pyrimidine nucleotides in a Zn2⫹- and Ni2⫹-dependent Met. Ions Life Sci. 2, 501–518 (2007)


513

manner. Unfortunately, the influence of nucleotides on the biochemical functions of SlyD has not yet been studied.

2.4.5.

SlyD Inhibits Mammalian Adenylate Cyclase

With the His6-tagged fragment of the intracellular domain of mammalian type 1 adenylate cyclase (IC1) from E. coli lysates, SlyD co-purified as a contaminant [23,30]. Among the individual isoforms of mammalian adenylate cyclase (AC1 to AC9) expressed in Sf9 insect cells, recombinant SlyD could inhibit the enzymatic activity of AC2 and AC7. In contrast, it increased the activity of AC5 and AC6. SlyD variants that display substantially reduced PPIase activity also show a diminished effect on adenylate cyclase activity [30]. SlyD-dependent regulation of adenylate cyclase activity might involve proline-mediated metastable folding states of the membrane-bound mammalian enzyme, as already discussed for the control of EGF-receptor autophosphorylation following extracellular EGFbinding by human FKBP12 [32].

3. 3.1.

INSIGHTS INTO THE BIOLOGICAL ROLE OF SlyD Lysis of E. coli by Bacteriophage ΦX174 Requires slyD

The multiplicity of FKBP genes in E. coli may reflect the fact that some of the FKBP have taken on specialized functions and are normally not involved in general catalysis of protein folding. The uncertainty about the biological roles of FKBP is in opposition to the ubiquity and high redundancy of the FKBP proteins. SlyD is a notable exception in this regard, because a strict phenotype could be observed for a slyD⫺ E. coli strain. It appeared that the host protein SlyD is essential for phage protein E mediated lysis of bacteriophage ΦX174 infected E. coli cells [22,33–36]. The lysis gene E of ΦX174 encodes a 92 amino acid membrane protein. Despite the multiplicity of PPIases in E. coli, the effect of protein E on lysis relies mostly on SlyD [37,38]. Only a partial compensation of lysis-inactivating slyD mutations was obtained by overexpressing two E. coli cyclophilins [38]. Wild-type ΦX174 phages accumulate to abnormally high levels in the slyD⫺ host strain, just as ΦX174-E⫺ phages do in wild-type E. coli. Western blots and pulse-chase labelling experiments showed that protein E is highly unstable in a slyD⫺ background, perhaps due to a lack of SlyD-catalyzed interconversion of a protease-sensitive native state isomer of protein E. The lysis factor E contains five proline residues, including three in the functionally important N-terminal part of the protein. A protein E mutation bypassing the SlyD requirement for Met. Ions Life Sci. 2, 501–518 (2007)

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lysis possesses the Leu19Phe substitution that is adjacent to the critical Pro21 [38]. An alternative explanation could be that protein E binds and alters the SlyD activity needed for some crucial step in maintaining envelope integrity. Thus, protein E may function through SlyD by disturbing the normal process of septation, because overexpression of E leads to extended macroscopic septal or polar lesions [39]. It is noteworthy that in the phage fd the cis/trans isomerization of proline213 of the phage 3 protein directly controls the E. coli-directed infectivity of the filamentous phage [40]. It is still unknown whether SlyD contributes to infectivity in this case. However, a FKBP-like PPIase was identified among the host cell proteins modified by phage T4 mono-ADP-ribosyltransferases in the course of the T4 replication [41].

3.2.

SlyD is Linked to the Hydrogenase Biosynthetic Pathway

Hydrogenases are enzymes that catalyze the reversible reduction of protons yielding dihydrogen (H2) as a reaction product. E. coli hydrogenase enzymes contain iron–sulfur clusters and two metal atoms (Ni and Fe) at their active site forming the [NiFe]-hydrogenases. The formation and insertion of the metal atoms into the metallocenter of [NiFe]-hydrogenases is a very complex process, in which auxiliary proteins, for example the products of the hypA-G genes, are involved [42]. Using the sequential peptide affinity system, SlyD was identified as a component of the HypB multiprotein complex [43]. To test whether SlyD has an important function in the hydrogenase biosynthetic pathway, a slyD deletion mutant was constructed. In fact, deletion of the slyD gene resulted in a significant reduction of hydrogenase activity in cell extracts prepared from anaerobic cultures. This phenotype could be rescued by nickel supplementation of the growth medium. Moreover, in vivo experiments with radioactive nickel have shown that slyD⫺ cells accumulate less nickel in comparison to the wild-type and that overexpression of SlyD can double the level of cellular nickel [43]. The result indicated that SlyD involvement relied more on nickel insertion than on a preceding enzyme maturation step. However, the exact mode of action is not yet clear. It is possible that SlyD affects the nickel uptake or export mechanisms of E. coli and thus indirectly influences hydrogenase production, but the physical interaction with HypB suggests that SlyD is more directly involved. Additionally, metal-binding histidine-rich stretches, like the CTE of SlyD, are observed in accessory proteins of biosynthetic pathways for other nickel-containing enzymes, such as urease and carbon monoxide dehydrogenase [44]. These stretches appear to be for nickel storage only and can be separated clearly from the nickel insertion activity. Therefore, SlyD is also a potential nickel source for the hydrogenase metallocenter assembly pathway and seems to be implicated in common nickel homeostasis of E. coli.

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CONCLUSIONS

Prolyl cis/trans isomerization forms a clever on/off switch for folding of the protein backbone. It is utilized for dynamic control of the physiological function of a protein in its various folding states. The energetic barrier of the prolyl bond switch is subject to regulated decrease by enzymes, the PPIases, in a spatial/temporal manner. The bacterial FKBP family of PPIases harbors SlyD that forms the unique example of a transition metal ion-regulated folding helper enzyme. The metal ion-binding site and the PPIase site were shown to be located on two different domains that communicate with each other in the Ni2⫹-mediated inhibition of the PPIase activity. It will be fascinating to follow the elucidation of the biological role of SlyD in phage replication and the bioformation of functional hydrogenases. However, it must be noted that no cellular SlyD protein substrate has been identified to date. For a given PPIase, polypeptide substrates might present many interaction sites in the respective Michaelis complexes, the number of which depends on the nature of the enzyme. Presumably, SlyD belongs to the PPIases which utilize a large number of secondary binding sites for efficient catalysis. Thus, the pronounced holding function of SlyD for unfolded polypeptide chains is an auxiliary factor to the PPIase activity, though remarkable from a biotechnological point of view. Understanding of this biological role requires the isolation and characterization of SlyD enzymes from other species, including those which do not contain FKBPs other than SlyD in the genome. Finally, the interplay between the metal ion-binding domain and the PPIase domain in the inhibitory response to Ni2⫹ ions can only be elucidated on the basis of a crystal or NMR structure of SlyD.

ABBREVIATIONS AND DEFINITIONS ADP ATP CD CsA CTE Cyp E. coli Ec EGF FKBP GTP H. influenzae

adenosine 5′-diphosphate adenosine 5′-triphosphate circular dichroism cyclosporin A C-terminal extension (amino acids 142 to 196 of SlyD) cyclophilin Escherichia coli prefix for Escherichia coli epidermal growth factor FK506-binding protein guanosine 5′-triphosphate Haemophilus influenzae

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HypB IC1 IF IMAP kcat kcat /Km KD Ki Km Mip Mt Par PPIase SlyD slyD⫺ Suc SurA UTP Xaa; Yaa ∆G‡

protein which was encoded by the hypB gene part of intracellular domain of mammalian type 1 adenylate cyclase insertion into flap segment immobilized metal affinity partitioning turnover number specificity constant dissociation constant inhibition constant Michaelis–Menten constant macrophage infecticity potentiator of Legionella pneumophila prefix for Methanococcus thermolithotrophicus parvulin peptidyl prolyl cis/trans isomerase EcFKBP21; sensitivity to lysis denotes a deletion mutant of the gene succinyl survival protein A uridine 5′-triphosphate any proteinogenic amino acid residue free energy of activation

Prolyl bond refers to the imidic peptide bond preceding proline in an amino acid sequence. Prolyl isomerization refers to the cis/trans isomerization of the peptide bond preceding proline.

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14 Chaperones of Nickel Metabolism Soledad Quiroz,1 Jong K. Kim, 2 Scott B. Mulrooney, 3 and Robert P. Hausinger1,2,3 1

Department of Biochemistry and Molecular Biology, 2 Cell and Molecular Biology Program, and 3 Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI 48824, USA ⬍[email protected]⬎ ⬍[email protected]⬎ ⬍[email protected]⬎ ⬍[email protected]⬎

1.

INTRODUCTION TO NICKEL METABOLISM 1.1. Nickel-Containing Enzymes 1.1.1. Urease 1.1.2. Hydrogenase 1.1.3. Carbon Monoxide Dehydrogenase 1.1.4. Acetyl-Coenzyme A Synthase/Carbon Monoxide Dehydrogenase 1.1.5. Methyl Coenzyme M Reductase 1.1.6. Superoxide Dismutase 1.1.7. Glyoxalase 1.1.8. Aci-reductone Dioxygenase 1.2. General Features of Nickel Incorporation into Proteins 1.2.1. Transport of Nickel into the Cell 1.2.2. Additional Processing of Nickel in the Cell 2. NICKEL METALLOCHAPERONES 2.1. Urease Metallochaperone: UreE 2.2. Hydrogenase Metallochaperones: HypA, HypB, and SlyD



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2.3. Carbon Monoxide Dehydrogenase Metallochaperone: CooJ 2.4. Other Potential Metallochaperones 3. MOLECULAR CHAPERONES INVOLVED IN NICKEL METABOLISM 3.1. Urease Molecular Chaperones: UreD, UreF and UreG 3.2. Hydrogenase Molecular Chaperones: HypC and HypB 3.3. Carbon Monoxide Dehydrogenase and Acetyl-Coenzyme A Synthase Molecular Chaperones: CooC/AcsF 4. CONCLUSIONS AND REMAINING QUESTIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

1.

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INTRODUCTION TO NICKEL METABOLISM

Nickel is an essential micronutrient of many organisms where it serves as a cofactor for enzymes involved in several critical metabolic processes [1,2]. Like other transition metal ions, however, excess Ni is toxic to cells; thus, synthesis of these Nienzymes requires the presence of carefully controlled Ni-processing mechanisms that range from selective transport of Ni into the cells to productive insertion of Ni into the apoproteins. Various accessory proteins participate in these processes and are required for the biosynthesis of several Ni-dependent enzymes. Here, we provide brief overviews of the catalytic activities, biological roles, and active site architectures of eight structurally characterized Ni-dependent enzymes. In addition, we summarize what is known about activation of these Ni-enzymes and emphasize two particular accessory protein roles: metallochaperones that bind and deliver Ni to the apoprotein forms of the enzymes and molecular chaperones that ensure productive conformations of the apoproteins for Ni incorporation.

1.1. 1.1.1.

Nickel-Containing Enzymes Urease

Urease (described further in Chapter 6) catalyzes the hydrolysis of urea to produce ammonia and carbamate. The latter molecule spontaneously decomposes to yield another molecule of ammonia and carbonic acid (Equations 1 and 2). Met. Ions Life Sci. 2, 519–544 (2007)

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This enzyme, found in plants, fungi and bacteria, has several biological roles, including its participation in recycling of environmental nitrogen and its use as a virulence factor in pathogenic microorganisms that are associated with gastric ulceration and urinary stone formation [3]. H 2 N-CO-NH 2 ⫹ H 2 O → H 2 N-COOH ⫹ NH 3

(1)

H 2 N-COOH ⫹ H 2 O → H 2 CO3 ⫹ NH 3

(2)

Crystallographic analyses have revealed that most bacterial ureases possess three structural subunits (encoded by ureA, ureB, and ureC) associated into a trimer of trimers [(αβγ)3], with each UreC subunit containing a dinuclear Ni active site bridged by a carbamylated Lys residue [4–6] (Figure 1A). Some species, such as Helicobacter pylori, have only two subunits (UreA, corresponding to a fusion of the small subunits in other bacteria, and the large subunit, labeled UreB) in a (α3β3) 4 supramolecular structure [7]. Plants and fungi have a single subunit corresponding to a fusion of all of the bacterial subunits, and form an α6 urease structure [8] that resembles a dimer of the bacterial enzyme. The urease gene cluster of most bacteria is composed of both structural genes (ureABC) and accessory genes (typically including ureDEFG, with additional urease-related genes present in some species). The structural gene products assemble into an apoprotein that requires activation by the accessory proteins. The best-studied urease activation system is that found in Klebsiella aerogenes, which contains the ureDABCEFG gene cluster [9,10]. Using this system, UreD, UreF, and UreG were identified as forming a GTP-dependent molecular chaperone that binds urease apoprotein [11], while UreE was shown to function as a metallochaperone that delivers Ni [12,13]. The detailed process of urease activation will be reviewed in Sections 2.1 and 3.1.

1.1.2.

Hydrogenase

Hydrogenases (described further in Chapter 7 of this volume) catalyze the reversible oxidation of molecular hydrogen into protons and electrons (Equation 3). These enzymes provide a mechanism for many microorganisms to use H2 as an energy source by generating a proton gradient or to remove excess reducing power in the form of molecular hydrogen [14]. H2 2 H+ + 2 e−

(3)

Three distinct classes of hydrogenases are defined by the metal content of their active sites: [NiFe]-hydrogenases, [Fe]-hydrogenases, and [iron-sulfur-clusterfree]-hydrogenases [14,15]. The crystal structures of several [NiFe]-hydrogenases have been resolved, including those of Desulfovibrio gigas and Desulfomicrobium baculatum [16–18]. Each heterodimeric protein has three iron–sulfur clusters in Met. Ions Life Sci. 2, 519–544 (2007)

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their small subunit and a [NiFe] active site in their large subunit. The active center contains Ni coordinated by four Cys residues (or three Cys and a selenocysteine in the D. baculatum enzyme), two of which bridge to the Fe that is also liganded by one carbon monoxide and two cyanide groups (Figure 1B). Seven accessory proteins are required to synthesize the Escherichia coli HycGE NiFe-hydrogenase [19], the paradigm system for defining the activation process of these enzymes [20,21]. These accessory proteins are the products of the six hyp genes (hypABCDEF) and another gene encoding a specific endopeptidase (hycI). The current model of HycE (large subunit) maturation includes a complicated series of steps involving: (1) HypDEF-mediated formation of an Fe(CN)2(CO) site in a process facilitated by HypC [22]; (2) insertion of Fe and its ligands into the precursor of the large subunit (retaining its C-terminal extension) when in complex with HypC [23]; (3) GTP-dependent addition of Ni to the active center mediated by HypAB; and (4) proteolytic processing of the C-terminus of HycE by HybD, leading to internalization of the catalytic center. Carbamoylphosphate (CP) was shown to be the precursor of the CN ligands [24], but CO is generated using a distinct substrate [25]. HypF displays CP phosphatase activity and catalyzes a CP-dependent pyrophosphate ATP exchange reaction [26]. More importantly, HypF catalyzes the ATP-dependent transfer of the carbamoyl group of CP to the C-terminal Cys of HypE [27]. The ATP-dependent dehydration of the thiocarbamoyl moiety by HypE results in thiocyanated HypE which can provide the CN ligand to Fe [27]. HypC is a central protein in the metallocenter assembly process [22,23], and is presumed to serve as a molecular chaperone (Section 3.2). HypA and HypB together appear to function as the Ni metallochaperone (Section 2.2), with the latter protein also suggested to have a molecular chaperone role (Section 3.2). The HycI endopeptidase subsequently cleaves off the C-terminal extension of HycE after Ni insertion, and Figure 1. Active sites of structurally characterized Ni-containing enzymes. In each case, Ni is shown as a solid black sphere and other elements are shown in various shades of gray. (A) Dinuclear Ni–Ni active site of urease (PDB code 1FWJ). The metal-bridging side chain is a carbamylated Lys and the three spheres coordinated to the metals are water molecules. (B) Dinuclear Ni–Fe active site of a [NiFe] hydrogenase (PDB code 1CC1). The Ni is bound to a seleno-Cys and three Cys (or to four Cys in related enzymes), two of which also coordinate the Fe. The Fe-bound diatomic ligands are two cyanide and one carbon monoxide molecules. (C) [Ni–Fe4 –S5] cluster of CODH (PDB code 1SU8). The structure of this cluster slightly varies in other CODH sites. (D) [4Fe–4S]–Ni–Ni site of ACS (PDB code 1RU3). The fourth ligand on the central Ni is water. (E) F430 Nitetrapyrrole of methyl coenzyme M reductase (PDB code 1MRO). (F) The active site of Ni-SOD (PDB code 1T6U). His1 is a ligand in the active enzyme, when the Ni is oxidized. (G) Ni-glyoxylase showing two bound water molecules (PDB code 1F9Z). His5 and Glu56 are derived from one subunit and His74 and Glu122 from the second subunit in the symmetric dimer. The two water molecules are displaced by substrate. (H) Ni-containing form of ARD as derived by a combination of solution structure analysis and homology modeling (PDB code 1M4O). The non-side-chain ligands of the metal are water molecules. Met. Ions Life Sci. 2, 519–544 (2007)

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the truncated protein binds to the small subunit and becomes membrane associated. HybD, a homolog of HycI that is specific to hydrogenase 2, has been structurally characterized and shown to possess a pentacoordinate Cd atom at the active site [28]. The apoprotein form of the endopeptidase is proposed to bind to hydrogenasebound Ni (coordinated by only three Cys ligands), thus activating the protease to cleave the correct peptide bond in the hydrogenase subunit [29–31].

1.1.3.

Carbon Monoxide Dehydrogenase

Carbon monoxide dehydrogenases (CODHs) (discussed further in Chapter 9) catalyze the reversible oxidation of carbon monoxide to carbon dioxide (Equation 4). Organisms possessing these enzymes play critical roles in the global carbon cycle and the degradation of environmental pollutants [32]. CO ⫹ H 2 O CO2 ⫹ 2 H⫹ ⫹ 2 e⫺

(4)

Crystal structures are known for CODHs from Carboxydothermus hydrogenoformans and Rhodospirillum rubrum [33,34]. Both proteins are ⬃130-kDa homodimers containing five metal–sulfur clusters of three types (B, C, and D) in a C-B′-D-B-C′ arrangement where the D cluster bridges the two subunits. While the B, B′, and D sites are the same cubane type [4Fe-4S] clusters in both proteins, the structures of the active site clusters (C and C′) slightly differ in the two proteins. The C cluster of R. rubrum is essentially a [1Ni–3Fe–4S] cubane bridged to a mononuclear Fe site, whereas the structure of the C. hydrogenoformans enzyme can be viewed as a [3Fe–4S] cluster fused with a [Ni–S–Fe] fragment containing a bridging sulfide (Figure 1C). Information regarding the mechanism of Ni insertion into CODH is available for R. rubrum where the cooCTJ gene cluster [35], located downstream of the cooS structural gene, is known to be involved. The CooC protein, which contains a nucleotide-binding motif, acts as an ATP/GTP-dependent molecular chaperone (see Section 3.3), while CooJ delivers Ni by using its histidine-rich C-terminal motif (Section 2.3).

1.1.4.

Acetyl-Coenzyme A Synthase/Carbon Monoxide Dehydrogenase

The CODH activity described above is found in another set of enzymes isolated from acetogenic bacteria and methanogenic archaea. The acetyl-CoA synthase/ carbon monoxide dehydrogenases (ACS/CODHs) (also discussed in Chapter 9) are bifunctional catalysts that exhibit the activity shown in Equation (4) and additionally synthesize (or decompose) acetyl-coenzyme A (CoA-SH) using the remarkable chemistry shown in Equation (5). The CODH site of ACS/CODH reduces CO2 to CO and then this gaseous molecule traverses a molecular tunnel within the protein to reach the ACS site where it is joined to CoA-SH and the Met. Ions Life Sci. 2, 519–544 (2007)


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methyl group from the corrinoid–iron–sulfur protein (Co-FeSP). Along with the monofunctional CODHs, these enzymes play a major role in the global carbon cycle and in the formation and removal of greenhouse gases [36]. CO ⫹ CoA-SH ⫹ CH 3 -Co(III)-FeSP CH 3 C(O)-S-CoA ⫹ Co(I)-FeSP (5) Crystallographic studies of Moorella thermoacetica ACS/CODH revealed that the tetrameric protein contains the dimeric CODH subunits at its core and one ACS subunit on each end [37,38]. The ACS metallocenter is a [4Fe-4S]-Ni-Ni site called the A-cluster. The [4Fe-4S] cluster is bridged to one Ni via a Cys side chain, and this metal is in turn bridged by two Cys residues to a second Ni, that is additionally bound by two backbone amides. The central Ni in the A-cluster is subject to metal substitution, resulting in inactive Cu–Ni and Zn–Ni species that were critical to identifying closed and open conformations of the protein. The [4Fe–4S]–Ni–Ni cluster (Figure 1D) also was observed in the structure of the monomeric C. hydrogenoformans ACS [39]. Little is known about the mechanism concerning metallocenter assembly in ACS/CODHs. Since the enzyme has two different sets of Ni-containing active sites, it is anticipated that several accessory proteins are required for biosynthesis. Consistent with this notion, ACS/CODH gene clusters contain several nonsubunit open reading frames (ORFs). In particular, AcsF encodes a CooC-like protein that is further described in Section 3.3.

1.1.5.

Methyl Coenzyme M Reductase

Methyl coenzyme M reductase (detailed further in Chapter 8) catalyzes the reaction of methyl-S-coenzyme M (CH3-S-CoM, where CoM-SH is 2-thioethanesulfonate) with coenzyme B (CoB-SH, N-7-mercaptoheptanoylthreonine phosphate) to form methane and the heterodisulfide, CoM-S-S-CoB (Equation 6). This is the final step of methane formation in methanogenic archaea growing on simple molecules such as acetate, methanol, formate, and carbon dioxide plus hydrogen gas [40]. CH 3 -S-CoM ⫹ CoB-SH → CH 4 ⫹ CoB-S-S-CoM

(6)

The X-ray crystal structure of methyl coenzyme M reductase, first obtained from Methanothermobacter marburgensis [41], reveals that the protein is a 300-kDa heterohexamer of three different subunits (α2β2γ2) containing two molecules of the Ni-containing tetrapyrrole, F430 (Figure 1E). This cofactor, named on the basis of its characteristic absorbance maximum at 430 nm when in the Ni(II) state, must be in the Ni(I) state for the enzyme to be active. Each active site F430 is buried deep in the protein and accessible from the surface by a 50-Å-long channel composed of mainly hydrophobic amino acids through which CH3-S-CoM Met. Ions Life Sci. 2, 519–544 (2007)

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can enter, and which is blocked by the binding of CoB-SH. An interesting aspect of this enzyme is the presence of five post-translationally modified amino acids near the active site: thio-Gly, N-methyl-His, S-methyl-Cys, 5-methyl-Arg, and 2methyl-Gln. Labeling studies have shown that the methyl groups are derived by methyl group transfer from S-adenosylmethionine, and not from the methyl group of CH3-S-CoM. The biosynthetic pathway of F430 is an offshoot of those for other biological tetrapyrroles [42]. Early labeling studies demonstrated that F430 is derived from dihydrosirohydrochlorin, which is also the precursor of siroheme and corrinoids. The dihydrosirohydrochlorin is formed from 5-aminolevulinic acid via uroporphyrinogen III. The conversion of dihydrosirohydrochlorin to F430 requires several steps, including amidation of acetate groups on two rings, reduction of two double bonds, cyclization of an acetamide to form the five-membered ring, cyclization of a propionic acid to form the six-membered ring, and insertion of Ni. However, the order of these steps and the mechanism underlying the Ni insertion and F430 incorporation into the protein remain unknown. An enzyme of related interest is found in methanotrophic archaea [43], such as those located in microbial mats that catalyze the anaerobic oxidation of methane [44]. These prokaryotes, closely related to methanogens in the order Methanosarcinales, contain homologs of genes encoding methyl coenzyme M reductase [45] and possess an F430-like molecule with a 46 Da mass increase [44]. The mechanism by which these microbes essentially reverse the last step of methanogenesis remains unclear.

1.1.6.

Superoxide Dismutase

Superoxide dismutases (SODs) are ubiquitous metalloenzymes that function to protect biological molecules from oxidative damage by catalyzing the dismutation of superoxide anion radicals to peroxide and molecular oxygen (Equation 7). In addition to the well-known Cu,Zn-, Fe-, and Mn-containing SODs, recent studies have revealed the existence of Ni-SODs (the focus of Chapter 10) in Streptomyces species and some cyanobacteria. 2 O⫺2 ⫹ 2 H⫹ → H 2 O2 ⫹ O2

(7)

Crystal structures of Ni-SODs have been solved for S. seoulensis and S. coelicolor enzymes [46,47]. The proteins are homohexamers consisting of fourhelix bundle subunits. The N-terminal loop coordinates the active site Ni(III) in square pyramidal geometry using two thiolate side chains (Cys-2 and Cys-6), two backbone amides (His-1 and Cys-2), and the His-1 side chain ligand at the axial position. The axial ligand is lost in the reduced state, with Ni(II) becoming square planar (Figure 1F). Apoprotein structures show that the residues involved in binding Ni are disordered. Met. Ions Life Sci. 2, 519–544 (2007)


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Ni-SODs in Streptomyces species are products of sodN, which encodes a preprotein with an N-terminal extension of 14 amino acids. During SOD maturation, proteolytic cleavage precedes Ni binding and results in the creation of the six-residue Ni-binding site. Recently, ORFs with significant homology to Ni-SODs were identified in the genomes of several cyanobacteria including Prochlorococcus marinus MIT9313 [48]. In this microbe, an ORF located downstream of sodN and named sodX was suggested to be the peptidase for maturation of the Ni-SOD. Coexpression of sodX and sodN in an oxygen-sensitive E. coli strain restored oxygen tolerance in a Ni-dependent manner, indicating the production of a catalytically active enzyme and providing confirmatory evidence for the importance of SodX in Ni-SOD maturation. Ni-SOD activity in S. seoulensis is stimulated by the overproduction of CbiXhp, a Ni-binding protein, suggesting that it too may function in metallocenter assembly [49]. Contrary to this notion, the gene encoding CbiXhp is located between two genes suggested to function in cobalamin biosynthesis. Further studies are needed to elucidate the detailed maturation steps of Ni-SOD activation, including the mechanism of Ni incorporation to the enzyme.

1.1.7.

Glyoxylase

Glyoxylase I (Glx I, the topic of Chapter 11 of this volume) is the first of two enzymes in the pathway to convert cytotoxic methylglyoxal into non-toxic α-hydroxycarboxylic acids. It converts the hemimercaptal substrate, formed nonenzymatically from methylglyoxal and glutathione (GSH, Equation 8), to non-toxic S-D-lactoylglutathione (Equation 9), which is the substrate for Glx II (Equation 10). These enzymes are important for cellular protection because methylglyoxal can exert toxic effects by reacting with DNA, RNA and proteins. CH 3 -CO-CHO ⫹ GSH CH 3 -CO-C(OH)-SG

(8)

CH 3 -CO-C(OH)-SG → CH 3 -CH(OH)-CO-SG

(9)

CH 3 -CH(OH)-CO-SG ⫹ H 2 O → CH 3 -CH(OH)-COOH ⫹ GSH

(10)

Unlike the case for Glx I of humans, Saccharomyces cerevisiae, and Pseudomonas putida, where the active site metal is zinc, Glx I from E. coli is completely inactive in the presence of Zn and is maximally active with Ni [50]. Reduced activity is found in the enzyme substituted with Co, Cd, and Mn. Crystallographic analyses revealed that catalytically active forms of E. coli Glx I with Ni, Co, and Cd each have octahedral metal coordination (Figure 1G), which is also observed in the Zn-containing human enzyme, whereas the inactive Zncontaining E. coli protein displays a five-coordinate metal site [51]. Several other pathogenic microorganisms are hypothesized to possess a Ni-containing Met. Ions Life Sci. 2, 519–544 (2007)

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glyoxylase on the basis of sequence comparisons [52]. The cellular mechanism of Ni incorporation into Glx I is unknown.

1.1.8. Aci-reductone Dioxygenase (ARD) Many microorganisms utilize the methionine salvage pathway to regenerate methionine from methylthioadenosine, produced during polyamine biosynthesis from S-adenosylmethionine. Aci-reductone is a key intermediate of this pathway, and is oxidized to two different sets of products in Klebsiella pneumoniae. One oxidation pathway leads to production of formate and the ketoacid precursor of methionine. The other route of oxidation, a non-productive pathway, converts the aci-reductone to formate, carbon monoxide, and methylthiobutyric acid. Remarkably, the two reactions are carried out by the same enzyme, ARD (described further in Chapter 12), depending on which metal is bound at the active site (Fe or Ni, respectively) [53]. The solution structure of K. pneumoniae Ni-ARD was determined by NMR methods [54]. The enzyme is a monomer containing two β -sheets that hinge together to form a ‘jelly roll’. Unfortunately, paramagnetism of the bound Ni causes broadening of the 1H resonance lines from residues near the metal center, thus hindering the structural characterization of the active site. Biophysical studies suggest the presence of three His ligands to the Ni, along with three other nitrogen or oxygen atoms [55]. Homology modeling of the active center, based on the structure of jack bean canavalin (another member of the cupin family), provides a reasonable model of the active site (Figure 1H). The mechanism of Ni insertion into the enzyme is unknown.

1.2. General Features of Nickel Incorporation into Proteins 1.2.1.

Transport of Nickel into the Cell

The first required step for synthesis of any Ni-enzyme is for the cell to take up the metal ion from the environment in a regulated manner. Bacteria have developed two major types of high-affinity Ni transport systems for efficient Ni uptake [56,57]: ABC-type transporters and Ni-specific permeases. The best-characterized ABC-type transporter system is that encoded by the E. coli nikABCDE operon [58], which is regulated by the product of a downstream gene, nikR. NikA is a periplasmic Ni-binding receptor protein with a Ni dissociation constant (Kd) of less than 0.1 µM (ten-fold lower than for Co, Cu, or Fe) [59]. Two crystal structures of metal-bound NikA reveal pronounced differences at the metal-binding site: in one case, a penta-hydrated Ni ion is suggested to be bound (with a single polar interaction) within a large cavity of the protein [60], whereas Met. Ions Life Sci. 2, 519–544 (2007)


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the second study reported the binding of a monohydrated Fe-EDTA complex at the same site of the protein using many specific interactions between the chelator and the protein side chains [61]. The long (2.7 Å) average Ni–O bond distance of the first crystal structure is inconsistent with spectroscopic data suggesting a much shorter distance (2.06 Å) [62], whereas the latter structural data more easily accommodate this distance. It is likely that a yet unidentified chelated form of Ni, rather than the ion itself, is the physiological species recognized by NikA. NikB and NikC are hydrophobic transmembrane proteins forming a pore for passage of the metal. NikD and NikE bind and hydrolyze ATP, and couple this energy release to the transport process. Ni homeostasis is achieved by use of NikR, a Ni-specific transcriptional repressor that binds to the NikR box in the promoter of the nik operon in the presence of Ni, resulting in suppression of Ni uptake [63]. The structure of the E. coli NikR apoprotein reveals that the protein contains a tetramerization domain and a ribbon–helix–helix DNA binding domain [64]. That same study showed four Ni sites within the isolated tetramerization domain, and a separate study of intact Pyrococcus horikoshi NikR identified the same four sites as well as a low-affinity Ni-binding site in the protein [65]. The first Ni-specific permease (encoded by hoxN) was identified in Ralstonia eutropha [66]. HoxN is an integral membrane protein containing eight membranespanning segments according to membrane topology analyses, and is the prototype of a novel family of transition metal permeases. Transport assays showed that HoxN has a high affinity for Ni with a transport constant (Kt) of ⬃ 20 nM, but with very low capacity. HoxN homologs have been reported in many bacteria (e.g., HupN in Bradyrhizobium japonicum, NixA in H. pylori, NicT in Mycobacterium tuberculosis, and NhlF in Rhodococcus rhodochrous) and Nic1p in the fission yeast, Schizosaccharomyces pombe [67]. The absence of HupN resulted in low levels of hydrogenase activity in B. japonicum under Ni-limiting conditions. Nic1p and NixA are essential for urease activity, and NixA exhibits a Ni Kt of 11 nM. NhlF was originally identified as a Co transporter in R. rhodochrous J1, but subsequent reinvestigation revealed that the permease transports both Ni and Co with a slight preference for Co. Regulation of the genes encoding these permeases is not well studied.

1.2.2.

Additional Processing of Nickel in the Cell

Once Ni enters the cell, it must be delivered and incorporated into the correct binding sites of Ni enzymes. As summarized above, this process may require metallochaperones (described further in Section 2), molecular chaperones (discussed in Section 3), and a variety of other assembly steps. For example, apoprotein proteolysis is associated with synthesis of hydrogenases and Ni-SOD. Cofactor synthesis is required prior to incorporation of the F430 tetrapyrrole into methyl coenzyme M reductase. Finally, many enzymes require the incorporation Met. Ions Life Sci. 2, 519–544 (2007)

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of another constituent prior to addition of Ni, such as the lysine carbamate of urease, the Fe(CN)2 (CO) site of hydrogenase, and the iron–sulfur cluster components of CODH and ACS.

2.

NICKEL METALLOCHAPERONES

The term metallochaperone refers to a protein that reversibly binds a metal ion, transports it within the cell, and provides it for metallocenter assembly to the target apoprotein. A generalized mechanism of such a protein is depicted in Figure 2, but other proteins (such as molecular chaperones described in Section 3) may participate in the process.

2.1.

Urease Metallochaperone: UreE

Among the multiple accessory genes required for urease activation in most urealytic organisms, UreE appears to function as a metallochaperone that delivers Ni to the urease apoprotein. The first suggestion that UreE functions as a Nibinding protein was provided by the sequences of the K. aerogenes and Proteus mirabilis urease operons [9,68]. The carboxyl termini of these proteins contain His-rich regions consisting of 10 His in the last 15 residues for K. aerogenes and 9 of the last 10 residues in the case of P. mirabilis, indicative of a potential metalbinding role. Subsequent sequences of ureE genes from other sources reveal that the His-rich C-terminal region is common, but it is completely absent in some organisms [69]. Equilibrium dialysis studies of K. aerogenes UreE showed that about 6 Ni bind per dimeric protein [70], while metal-binding studies of Bacillus pasteurii UreE, which contains only two conserved His residues in this region, found a single Ni bound per dimer [71]. Purified UreE proteins also bind other metal ions, such as Cu and Zn, demonstrating that the specificity of urease for Ni does not reside solely with this delivery protein [72]. Using site-directed

Metallochaperone Holoprotein Ni

Ni-Metallochaperone

Figure 2.

Target Apoprotein

Generalized mechanism of a Ni metallochaperone.

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mutation methods to create a truncated form of K. aerogenes UreE with the last 15 amino acids removed (His144*UreE), the His-rich region was demonstrated to be nonessential; i.e., the truncated protein still binds 2-3 Ni ions per dimer and is still competent in facilitating Ni-dependent activation of urease in vivo [73,74]. In a complementary study, the native H. pylori ureE gene, which does not encode a protein with a His-rich C-terminal tail, was fused to several extensions to produce different His-rich regions [75]. The resulting His-rich variants show increased Ni-binding and cells containing these variants have increased urease levels; thus, the C-terminal His-rich region has a Ni-sequestering function that aids in urease activation. Several lines of evidence indicate that UreE interacts with other accessory proteins during Ni-dependent activation of urease. In vitro studies showed that a complex of urease apoprotein with UreD, UreF, and UreG is fully activated only by including UreE in a mixture containing GTP, bicarbonate, and Ni [13], thus providing strong evidence that UreE functions as a metallochaperone to deliver Ni to the UreDFG–urease apoprotein complex. These studies also showed that UreE does not simply function as a reversible Ni-binding protein because activation occurred, even when metal ion chelators were included in the reaction [13]. Additional work involving yeast two-hybrid analysis demonstrated an interaction between H. pylori UreE and UreG proteins [76,77]. Site-directed mutagenesis and structural studies provided detailed insights into the metal-binding properties of UreE. Variants of K. aerogenes His144*UreE affecting His-110 or His-112 exhibit reduced Ni binding while not greatly affecting urease activation, whereas a variant affecting His-96 binds less Ni and abolishes UreE’s capacity to activate urease [74]. These results are easily rationalized by the crystal structures of Cu-bound K. aerogenes H144*UreE [78] and Zn-bound B. pasteurii UreE [79]. The overall structures are nearly identical, but contain three and one metal-binding sites, respectively. Both proteins bind a metal at the dimer interface using the symmetric pair of critical His-96 residues in the K. aerogenes protein or the pair of His-100 residues for B. pasteurii UreE. This metal site is essential for UreE’s function in urease activation. In addition, each subunit of K. aerogenes UreE binds a metal at sites involving His-110 and His-112, residues that are substituted with other side chains in the B. pasteurii protein. The UreE crystal structures also reveal the presence of two distinct domains in the proteins. The metal-binding domains, located in the C-terminal half of each molecule, resemble the structure of the yeast copper metallochaperone Atx1 [80]. The N-terminal domains have structural similarities to a domain of the yeast Hsp40 molecular chaperone Sis1 [81], suggesting that this domain may be involved in molecular recognition and binding to other urease accessory proteins and/or urease apoprotein. Arguing against this conclusion are results from studies involving a construct that produced only the metal-binding domain of K. aerogenes UreE (residues 70-143); the single domain of UreE is capable of delivering Ni to the urease apoprotein and the N-terminal domain is not required [82]. Met. Ions Life Sci. 2, 519–544 (2007)

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Complicating the B. pasteurii UreE structure described above, the crystallization conditions promote oligomerization of the protein to form a dimer of dimers (α2)2 in which all four His-100 side chains serve as ligands to a single Zn [79]. The tendency of B. pasteurii UreE to aggregate was further examined by protein NMR and equilibrium dialysis approaches [83]. Those studies showed that the tetramer form is favored only at high protein concentrations and provided evidence for a second Ni-binding site in the C-terminal region of the dimeric form of UreE, which probably binds a total of 3 Ni ions. A final comment about urease metallochaperones focuses on the situation in H. pylori where HypA and HypB, normally associated with hydrogenase activation, are also required for urease activity. Deletions of either gene encoding these proteins results in cells with very low urease activity; however, the urease activity can be restored by addition of excess Ni [84,85]. Thus, it is possible that HypA, HypB, or a complex of these proteins function as a metallochaperone and assists in urease activation. These proteins are discussed further below.

2.2.

Hydrogenase Metallochaperones: HypA, HypB, and SlyD

Of the many proteins involved in maturation of [NiFe] hydrogenases, three are known to directly bind Ni and may function as metallochaperones: HypA, HypB, and SlyD. HypA-like and HypB-like proteins are given different designations depending on the organism and the particular hydrogenase system (see below). Also of note, both HypB and SlyD exhibit additional activities beyond their putative metallochaperone roles. Mutations in genes encoding any of these proteins result in hydrogenase deficiencies in vivo, and the activity can be partially restored by supplementation of the growth medium with Ni [19,86–90]. HypA designates the ⬃13-kDa protein required for activation of hydrogenase 3 of E. coli and the corresponding protein in many other hydrogenase systems. Homologs are termed HupA in selected microorganisms or HybF when referring to the protein used for E. coli hydrogenase systems 1 and 2. Purified HypA from H. pylori binds two Ni ions in a cooperative manner [85]. Site-directed mutagenesis studies revealed that a single His residue is required for binding Ni, and introduction of the corresponding mutant gene into the chromosome resulted in substantial loss of hydrogenase activity [85]. Subsequent investigations of E. coli HypA showed that it binds stoichiometric amounts of Ni and Zn, with µM and nM affinities, respectively [91]. Based on UV/visible spectroscopic results indicating thiolate ligation, the bound Zn is proposed to have a structural role. Similar mutagenesis and binding studies of E. coli HybF found a single histidine is necessary for Ni binding, but mutagenesis of this residue resulted in a protein that retained the ability to bind Zn [92]. Met. Ions Life Sci. 2, 519–544 (2007)


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HypB, alternatively termed HupB in certain microorganisms, is a ⬃30-kDa protein that contains a nucleotide-binding motif and possesses low levels of GTPase activity [85,86,93–95]. Site-directed mutations in the GTP-binding motifs result in elimination of hydrogenase activity [86], analogous to other molecular chaperones used for Ni-enzyme biosynthesis (see Section 3.2). In addition to its GTPase activity, HypB proteins of selected organisms contain His-rich regions that are capable of binding several Ni ions. For example, HypB from Rhizobium leguminosarum binds 4 equivalents of Ni [95] and B. japonicum HypB, which contains 24 His in a 39 residue region, binds 9 equivalents of Ni [94]. When this motif is deleted, the B. japonicum protein still binds one equivalent of Ni and retains competence in activating hydrogenase [94,96,97]. The best-studied HypB is that from E. coli. This protein lacks a His-rich region, yet it tightly binds one Ni per monomer (Kd of 0.12 pM) using a CXXCGC motif at the N-terminus [98]. Furthermore, the E. coli protein has a second metal-binding site located in the GTP-binding domain that has weaker affinity for either Ni or Zn ions [98]. In addition to the myriad studies of the individual HypA and HypB proteins and their homologs, there is significant evidence that the two proteins interact. Deficiencies of either HypA or HypB led to reduction of both urease and hydrogenase activities in H. pylori, suggesting an interaction between these two proteins [84,85]. Subsequent chemical cross-linking studies showed that a stable HypAHypB complex is formed for these proteins [85], as well as those from E. coli [91]. In contrast to these results, chemical cross-linking studies carried out with a Strep-tagged variant of HybF failed to detect an association with HypB [92]. Although the above studies of HypA and HypB proteins have greatly added to our knowledge, the question of how they function in Ni delivery and/or insertion into hydrogenase remains to be discovered. SlyD (see Chapter 13) is a ⬃21-kDa protein possessing an N-terminal region (146 amino acids with similarity to FK506-binding proteins) that contains peptidyl-prolyl cis/trans isomerase activity and a short C-terminal metal-binding region rich in His, Asp, Glu, and Cys [99,100]. Mutations in slyD result in cells that fail to lyse when infected with bacteriophage ΦX174, because SlyD stabilizes the E lysis protein [99,101]. Additional evidence suggests that SlyD may act as a chaperone for several other proteins [102–105]. More pertinent to this discussion, SlyD reversibly binds Ni in its C-terminal region such that the peptidyl/prolyl isomerase activity is inhibited [100]. The ability of SlyD to bind Ni was suggested by its co-purification with several recombinant His-tagged proteins when using Ninitrilotriacetic acid affinity chromatography [104,106–108]. Of particular interest with regard to maturation of the [NiFe] hydrogenases, E. coli SlyD was shown to interact with HypB from the same source [90]. Furthermore, cells deleted in slyD have greatly reduced activity of all three hydrogenases and reduced intracellular concentrations of Ni. These findings suggest that SlyD has a role in the Ni insertion step of hydrogenase activation [90]. Met. Ions Life Sci. 2, 519–544 (2007)

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2.3. Carbon Monoxide Dehydrogenase Metallochaperone: CooJ Two R. rubrum proteins are known to be involved in the insertion of Ni into the metallocenter of CODH. CooC is a membrane-associated protein that has ATPase and GTPase activity, but does not bind Ni, and is described as a putative molecular chaperone in Section 3.3. The second protein, CooJ, contains 115 residues of which 16 of the C-terminal 34 amino acids are His [109], an arrangement similar to the His-rich regions in sequences of some HypB and UreE proteins. Cells containing a chromosomal deletion that eliminates the His-rich region display Ni-dependence for growth on CO that is identical to wild-type strain, while cells with an insertional mutation of cooJ required 1000-fold higher Ni than wild type for optimal growth [35]. These findings suggest that the His-rich C-terminal region is not required for CooJ function, and the functional Ni-binding site lies elsewhere in the protein.

2.4.

Other Potential Metallochaperones

Several other proteins possess His-rich regions and/or tightly bind Ni; however, none of these has been convincingly shown to facilitate Ni metallocenter assembly. For example, CbiXhp of S. seoulensis contains a carboxyl terminus in which 11 of 19 residues are His, and the cellular overproduction of this protein stimulated Ni-SOD activity [49]. On the other hand, this protein is more likely to be involved in cobalamin biosynthesis on the basis of flanking genes and its presence in cells that lack a Ni-SOD. The Hpn protein of H. pylori is worth a few comments because of the high levels of Ni-containing urease and the important role of hydrogenase in this microorganism [110,111]. Hpn is comprised of only 60 amino acids, 28 of which are His [112]. Deletion of the corresponding gene has no effect on urease activity for cells grown on blood agar, but does make the cells more susceptible to growth inhibition at high Ni concentrations [113]. Recent studies have shown Hpn binds 5 Ni per monomer (Kd 7.1 µM) and provided evidence that the concentrations of this protein correlate to the intracellular Ni concentration and to the cell’s ability to tolerate high Ni concentrations [114]. The authors suggested that Hpn may function in Ni storage, Ni donation, and Ni detoxification. It will be of interest to learn whether follow-up studies confirm the putative metallochaperone role for this protein and to monitor whether Ni metallochaperones for other enzyme systems are identified.

3.

MOLECULAR CHAPERONES INVOLVED IN NICKEL METABOLISM

Molecular chaperones are proteins that prevent misfolding or assist in re-folding of other proteins, often by using energy derived from nucleotide hydrolysis. For Met. Ions Life Sci. 2, 519–544 (2007)


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Molecular Chaperone Holoprotein Target Apoprotein

(NTP) Ni (or Ni-Metallochaperone) Apoprotein: Molecular Chaperone

Figure 3. Generalized mechanism of a Ni-enzyme-specific molecular chaperone.

example, the best-studied molecular chaperones are the GroES:GroEL chaperonin, Hsp70, and Hsp40 that act on many cellular proteins [115,116]. Recent studies have shown that SlyD, a Ni-binding protein, similarly exhibits a diverse molecular chaperone role [102–105]. Such nonspecific molecular chaperones are likely to stimulate the proper folding of many Ni-containing enzymes, as evidenced by the diminished hydrogenase activity in groEL or groES mutants and by the specific binding of GroEL to the HycE precursor protein [117]. Indeed, such a role may explain some of the stimulatory effects of SlyD on E. coli hydrogenase activation [90]. While these housekeeping proteins are rather nonspecific in their action, this section focuses on putative molecular chaperones that are specific to individual Ni-enzyme activation systems. A generalized mechanism of a Ni-enzyme-specific molecular chaperone is depicted in Figure 3. As indicated, these proteins may drive the reaction by coupling metal insertion to nucleotide hydrolysis and/or they may use a metallochaperone rather than the free metal ion. In general, such proteins appear to be more essential to the activation processes than the metallochaperones, whose absence often can be overcome by excess Ni.

3.1.

Urease Molecular Chaperones: UreD, UreF and UreG

Structural studies have revealed that the Ni active site of urease is buried within the enzyme [4], and this site is also relatively inaccessible in the apoprotein [118]. These results are consistent with the need for one or more urease-specific molecular chaperone(s) to alter the urease protein conformation and allow Ni to gain access to the active site. From studies with the K. aerogenes urease system, three proteins, each of which is required for in vivo enzyme activation, are proposed [1,10] to act together to fulfill this role: UreD, UreF and UreG. As expected of molecular chaperone proteins, each of the UreD, UreF, and UreG accessory proteins are found in complexes that include the urease apoprotein. Thus, UreD-, UreDF-, and UreDFG-urease apoprotein complexes have been described [119-121]. These complexes possess distinct properties when compared Met. Ions Life Sci. 2, 519–544 (2007)

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with those of the urease apoprotein alone, especially with regard to their activation properties. Approximately 15% of the apoprotein is activated in vitro by addition of 100 µM Ni and 100 mM bicarbonate (needed to carbamylate the active site Lys) [122, 123]. In contrast, about 30% of the UreD-urease apoprotein is activated by these conditions, demonstrating that UreD directly enhances this process [119]. Furthermore, the UreDF-urease apoprotein is activated to the same extent by using nearly 1000-fold lower concentrations of bicarbonate, and activation of this complex is resistant to the detrimental effects of high concentrations of Ni compared with the apoprotein alone or to UreD-urease apoprotein [120]. A UreDFG-urease apoprotein complex forms upon addition of UreG to UreDFurease apoprotein [11] and is normally present in cells expressing the intact K. aerogenes urease gene cluster [121]. Significantly, this species exhibits GTPdependent urease activation—shown by mutagenesis studies to be associated with the nucleotide-binding (P-loop) motif located within UreG [11]. Urease activation is not achieved with a non-hydrolyzable analog of GTP. When the UreE metallochaperone is provided to this complex along with GTP and near-physiological levels of Ni plus bicarbonate, fully active urease is generated [13]. Three of the urease apoprotein species were probed by a chemical cross-linking/proteolysis/ mass spectrometric approach to examine the sites of binding of UreD and UreF to urease [124]. Additional evidence from these studies suggests that UreF interacts with UreDurease apoprotein to give rise to a conformational change within urease that may enhance access of Ni to its ligand-binding residues. Finally, we note that a UreDFG complex is generated in the absence of the structural proteins, and this species was enriched by binding to an ATP-linked agarose resin [125]. Further structural and mechanistic studies are required to better define the individual roles of UreD, UreF, and UreG within the heterotrimeric molecular chaperone that couples GTP hydrolysis to urease activation. In addition to participating as a component of the urease molecular chaperone, UreG has been studied in its purified form. The K. aerogenes protein is a monomer of 21.9 kDa that, despite having a P-loop motif, fails to bind or hydrolyze GTP [125]. In contrast, UreG from B. pasteurii, which is over 50% identical to the K. aerogenes protein, is dimeric and has weak GTPase activity [126]. NMR studies of the B. pasteurii protein suggest that it is intrinsically disordered; however, this UreG was purified from a heterologous overproduction system and required urea dissolution of inclusion bodies, so the disorder may be artifactual. Significantly, the purified B. pasteurii UreG binds 2 Zn per dimer (Kd 42 µM) or binds 4 Ni per dimer with weak affinity (Kd 360 µM). Two other potential molecular chaperones for urease exist in H. pylori. First, is the GroES homolog called HspA [127]. This heat shock protein possesses an N-terminal domain resembling the broad specificity molecular chaperones, but the protein additionally contains a 27 amino acid extension that is rich in His. Not unexpectedly, HspA binds 2 Ni with high specificity. Significantly, when Met. Ions Life Sci. 2, 519–544 (2007)


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hspA is co-expressed with the H. pylori urease genes in E. coli, urease activity is enhanced four-fold in accordance with a possible urease-specific function [127]. The second example of a possible H. pylori-specific molecular chaperone involves the typical hydrogenase accessory protein HypB which is required for both urease and hydrogenase activity in this organism. Since HypB is suggested to function as a molecular chaperone of hydrogenase (Section 3.2), it may also serve this role in urease activation in H. pylori. Alternatively, HypB along with HypA may possess a metallochaperone role in urease activation in this microorganism (Section 2.1).

3.2. Hydrogenase Molecular Chaperones: HypC and HypB The complex biosynthetic pathway of E. coli hydrogenase 3 (summarized in Section 1.1.2) includes two molecular chaperone-like proteins: HypC and HypB. E. coli contains a second HypC-like protein, termed HybG, which specifically binds to hydrogenase 2 (both HypC and HybG bind to hydrogenase 1, but only the former facilitates activation) [128]. Homologs to HypC and HypB are found in many other organisms where they are sometimes given alternative designations (e.g., HupB and HupC). The evidence supporting the roles of HypC and HypB as molecular chaperones is described below. HypC (or its homolog) plays a central and multifaceted role in [NiFe]hydrogenase biosynthesis. HypC forms a complex with HypD, especially when the synthesis of CP is reduced so that the formation of the Fe(CN)2 (CO) center is hindered [23]. The HypC-HypD species additionally binds HypE in such a manner as to allow its carbamoylation by HypF [22]. After dehydration of the modified HypE [27], the resulting cyano group is transferred to Fe within the HypC-HypD complex [22]. CO is presumably incorporated onto the Fe prior to or after this step, but this ligand is not derived from CP and a different, still uncharacterized, precursor is required [25]. HypD dissociates from HypC as a new complex, HypC–HycE, is formed [129,130]. The HypC–HycE complex contains the Fe(CN)2 (CO) center, formed in the earlier complex, but it still lacks Ni [129]. Formation of the HypC–HycE complex requires the presence of the hydrogenase subunit C-terminal 15 amino acid extension, as shown by the inability of a truncated variant to become activated. Furthermore, the interaction between the chaperone and the large subunit precursor requires the N-terminal Cys residue of HypC and a particular large subunit Cys residue, which eventually coordinates Ni in the active site [131]. Ni is provided by the action of HypA/HypB metallochaperone (Section 2.2) [87], with HypB additionally having a molecular chaperone role (described below). After the complete set of metallocenter components is in place, HypC dissociates, HycI binds to the HycE-bound Ni and becomes proteolytically active [29]. The large subunit extension is removed, resulting in the [NiFe] site becoming buried in the protein. Met. Ions Life Sci. 2, 519–544 (2007)

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Nickel insertion into the Fe(CN)2 (CO)-containing and HypC-bound hydrogenase precursor requires the HypA/HypB metallochaperone, with HypB also exhibiting a GTP-dependent molecular chaperone role [86]. HypB is homologous to the urease accessory protein UreG and likewise contains a nucleotide-binding motif. Native HypB exhibits weak GTPase activity, but a mutation affecting the P-loop eliminates the GTPase activity and greatly decreases the hydrogenase activity [84,85]. The HypB proteins of some organisms contain His-rich termini that are able to bind Ni; removal of this sequence has only a small effect on hydrogenase activity while having a larger affect on cellular Ni content. The HypB molecular chaperone appears to drive Ni insertion into the hydrogenase subunit by coupling this reaction to GTP hydrolysis [85,86,93–95].

3.3. Carbon Monoxide Dehydrogenase and Acetyl-Coenzyme A Synthase Molecular Chaperones: CooC/AcsF The CODH operon of R. rubrum encodes the suspected molecular chaperone CooC. This membrane-bound homodimer of 62 kDa is related in sequence to UreG of urease activation and HypB of hydrogenase biosynthesis [35]. CooC contains a P-loop motif in its N-terminus, and the purified protein hydrolyzes both GTP and ATP with similar Km values, but with a 10-fold greater Vmax for ATP. Mutation of residues in the P-loop motif prevents Ni insertion into CODH and abolishes the ATPase activity, both in vivo and in vitro. Ni is not present in the purified protein [132]. The gene cluster encoding the ACS/CODH bifunctional enzyme of M. thermoacetica includes a gene encoding AscF, that resembles CooC and the other nucleotide-dependent molecular chaperones described above. This protein contains a P-loop and has five conserved Cys in a motif characteristic of iron coordination. Despite the suspected importance of this gene for ACS/CODH activation, its deletion had no effect on enzyme activity. It remains possible that a second copy of the gene is present in the genome [133].

4.

CONCLUSIONS AND REMAINING QUESTIONS

Accessory proteins are clearly shown to play critical roles in the biosynthesis of several Ni-containing enzymes. As emphasized in this contribution, metallochaperones often are used to deliver Ni to the target apoprotein and nucleotidedependent molecular chaperones can have a role in altering the apoprotein conformation to allow access by the metal ion. Indeed, the two functions are usually closely linked. Even when such proteins have been identified, their precise mechanisms of action remain unclear. For example, it is unknown how Ni becomes bound to metallochaperones and how the Ni is delivered to the target Met. Ions Life Sci. 2, 519–544 (2007)


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protein. Similarly, the mechanism by which nucleotide hydrolysis is coupled to Ni insertion remains a mystery. Furthermore, these two themes are not universal among all Ni enzymes, as exemplified by Ni-dependent glyoxylase and aci-reductone dioxygenase, for which no accessory proteins have been observed; on the other hand, it remains possible that future investigations will reveal the existence of such accessory proteins for these cases. Finally, it is important to note that exceptions to the metallocenter assembly mechanism exist within the particular enzyme systems. For example, Bacillus subtilis synthesizes sufficient levels of Ni-containing urease to allow for growth on urea as sole nitrogen source, even though its genome lacks homologs to ureDEFG [134]. The mechanism by which this organism generates active urease in the apparent absence of any accessory protein remains unknown. Future biochemical investigations are certain to provide additional insights into the fascinating topic of Ni metallocenter assembly.

ACKNOWLEDGMENTS Our studies involving Ni metallochaperones, molecular chaperones, and other aspects of urease activation have been supported by the National Institutes of Health Grant DK45686 (to R.P.H.)

ABBREVIATIONS ACS ARD ATP CH3-S-CoM CoA-SH CoB-SH CODH Co-FeSP CoM-SH CP EDTA Glx GSH GTP NMR NTP ORF

acetyl-coenzyme A synthase aci-reductone dioxygenase adenosine 5′-triphosphate methyl-S-coenzyme M coenzyme A coenzyme B (N-7-mercaptoheptanoylthreonine phosphate) carbon monoxide dehydrogenase corrinoid–iron–sulfur protein coenzyme M (2-thioethanesulfonate) carbamoyl phosphate ethylenediamine-N,N,N′, N′-tetraacetate glyoxylase glutathione guanosine 5′-triphosphate nuclear magnetic resonance nucleoside 5′-triphosphate open reading frame Met. Ions Life Sci. 2, 519–544 (2007)

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PDB SOD

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protein data bank superoxide dismutase

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15 The Role of Nickel in Environmental Adaptation of the Gastric Pathogen Helicobacter pylori Florian D. Ernst,1 Arnoud H. M. van Vliet,1 Manfred Kist, 2 Johannes G. Kusters,1 and Stefan Bereswill*3 1

Department of Gastroenterology and Hepatology, Erasmus MC-University Medical Center, Rotterdam, The Netherlands

2 Department of Microbiology and Hygiene, Institute of Medical Microbiology and Hygiene, University Hospital of Freiburg, Freiburg, Germany 3

Charité - University Medicine Berlin, Institute of Microbiology and Hygiene, Campus Charité Mitte, Dorotheenstrasse 96, D-10117 Berlin, Germany ⬍[email protected]⬎

1.

INTRODUCTION 1.1. Microbiology and Molecular Biology of Helicobacter pylori 1.2. H. pylori Infection and Associated Diseases 1.2.1. Course of H. pylori Infection 1.2.2. Diagnostics 1.2.3. Treatment 1.3. Mechanisms of Pathogenesis 1.4. The Roles of Urease and Nickel in Gastric Adaptation 1.5. Mechanisms of Metal Ion Homeostasis 2. NICKEL ENZYMES AND ENVIRONMENTAL ADAPTATION 2.1. Urease 2.2. The Role of Urease in Acid Resistance



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2.3. Ni/Fe-Hydrogenase 2.4. Hydrogenase and Gastric Adaptation NICKEL UPTAKE SYSTEMS 3.1. Nickel Import by the NixA Protein 3.2. Possible Alternative Nickel Transporters MECHANISMS OF NICKEL REGULATION 4.1. The NikR Regulator 4.2. Genes Regulated by NikR 4.2.1. Regulation of Nickel Uptake 4.2.2. Control of Urease Synthesis 4.2.3. Modulation of Fur and Amidase Genes 4.2.4. Regulation of the NikR Gene 4.2.5. Hydrogenase and Other Genes 4.3. Metal Metabolism and Acid Regulation PROTECTION OF NICKEL METABOLISM 5.1. Nickel Binding and Storage 5.2. Specific Export of Other Metal Ions METAL METABOLISM AS DRUG TARGET: THERAPEUTIC CONSIDERATIONS 6.1. Vaccination 6.2. Inhibition of Metalloenzymes 6.3. Metal Diet CONCLUSIONS ABBREVIATIONS REFERENCES

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INTRODUCTION

Diseases of the upper intestinal tract affecting stomach and duodenum represent a common problem that nearly everybody encounters during life. Before the discovery of the gastric pathogen Helicobacter pylori, such disorders remained debilitating and incurable in many cases. The first report on spiral bacteria in the stomach of non-human mammals dates back over 100 years [1]. Early reports suggested a role for gastric infection by at that time unknown bacteria, and also a causal association between chronic bacterial antrum gastritis and duodenal ulcer, which was described in detail in 1923, by the German surgeon Konjetzny [2]. However, scientific proof for bacteria as causes for gastric diseases did not come until 1982, Met. Ions Life Sci. 2, 545–580 (2007)

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when Robin Warren and Barry Marshall, together with the local microbiologists, isolated H. pylori, originally named Campylobacter pyloridis, from gastric biopsy samples [3]. Subsequently, Marshall fulfilled Koch’s postulates for H. pylori in a selfinfection experiment, which resulted in H. pylori-induced gastritis. The bacteria were re-isolated from gastric biopsies, and inflammation was cured by subsequent antibiotic treatment [4]. The discovery of H. pylori infection as causative agent of gastric and duodenal disorders was a major breakthrough in gastroenterology, for which Marshall and Warren were awarded the Nobel Prize in medicine in 2005. This discovery offered not only a novel idea for the understanding of mechanisms underlying gastritis, ulcerations, lymphomas and gastric cancer, but also for the treatment of gastric diseases by antimicrobial therapy. In this review, we focus on the essential role of H. pylori nickel ion metabolism and metal ion homeostasis in gastric adaptation and highlight the extraordinary role of nickel in acid resistance required for both primary colonization and long-term survival in the hostile gastric niche.

1.1.

Microbiology and Molecular Biology of Helicobacter pylori

H. pylori belongs to the ε-subdivision of proteobacteria, and is classified in the Helicobacteraceae family of the Campylobacterales order [5]. It is a highly motile non-spore-forming, gram-negative, curved bacterium with an average size of 0.2–1.2 ⫻ 1.5-7 µm. Motility is mediated by 5–6 unipolar flagella, which are sheathed and end in a terminal bulb [6]. H. pylori requires microaerobic conditions for growth, and is usually cultivated in a nitrogen atmosphere containing 2–5% O2 and 5–10% CO2 [7]. The standard identification method for H. pylori in gastric biopsy material is Gram or Warthin-Starry (silver) staining followed by microscopy. Biochemical key reactions of species differentiation include tests for urease, catalase and oxidase, as H. pylori is positive for all three enzymes [7]. In 1997 and 1999 the genome sequences of two H. pylori strains (26695 [8] and J99 [9]) became available. The H. pylori genome has an average size of 1.6 Mbp with an overall G/C-content of 35%. Approximately 1600 open reading frames (ORF) were identified which cover 90% of the H. pylori genome. One-third of the ORFs are considered to be Helicobacter-specific [8–10]. About 50% of all H. pylori strains carry plasmids [11], which contribute to genetic diversity [12] and were successfully applied for genetic manipulation and cloning [13]. The H. pylori genome is highly diverse and individual strains adapt to their personal human host by point mutations, nucleotide substitutions, insertions and deletions [14–16]. In addition, H. pylori is able to switch transcription and translation of genes on and off through phase variation [17]. Sequence comparison of H. pylori gene fragments from isolates all over the world revealed that the bacteria populations have been colonizing humans for more than a hundred thousand years [18], exhibiting a high degree of co-evolution with the host. This was demonstrated by the fact that H. pylori populations arising in Africa and Asia were spread worldwide through human migratory fluxes [19]. Met. Ions Life Sci. 2, 545–580 (2007)

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1.2. H. pylori Infection and Associated Diseases H. pylori colonizes the gastric mucus overlaying the epithelial cells, and this results in lifelong chronic infection unless treated [16,20,21]. H. pylori infection occurs worldwide, and the prevalence varies between different geographic regions. Infection takes place in early childhood, by direct transmission, mostly from mother to child [22]. The incidence of infection is strongly associated with the socioeconomic status. In developing countries around 90% of the population is infected, and this is closely related to low income, high numbers of children per household, and low hygiene standards [22]. In contrast, the infection concerns only 20–50% of the population in industrialized countries. This most probably results from higher hygiene standards during childhood and to a lower extent to active antibiotic treatment. Over the last two decades the prevalence of H. pylori infection has decreased significantly in the Western world [16].

1.2.1.

Course of H. pylori Infection

It is generally accepted that H. pylori infection causes acute gastric inflammation followed by autoimmune reactions (Figure 1). Gastritis develops predominantly in the antral part of the normacidic or hyperacidic stomach, where the local acid concentration is lower compared with the gastric corpus [23]. There, it might further develop into gastric and duodenal ulcerations (Figure 1). In contrast, patients with low acid output show gastritis in the corpus predominantly, and are at increased risk of developing gastric ulcers, which can develop into atrophy, intestinal metaplasia, gastric carcinoma and MALT lymphoma [16]. As a consequence, H. pylori was classified a class I carcinogen by the WHO in 1994 [24]. Finally, H. pylori infection might influence gastroesophageal reflux

Metaplasia

Gastric Cancer MALT Lymphoma

Gastritis

Pepsin/ Acid

Asymptomatic Infection

Ulceration

Figure 1. Overview on H. pylori-associated diseases. The infection always leads to gastritis. The further outcome of infection can differ, as indicated by the arrows. Met. Ions Life Sci. 2, 545–580 (2007)


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disease (GERD), as GERD patients were shown to have a lower prevalence of H. pylori infections. It is thought that the bacteria might prevent GERD by suppression of the acid-producing parietal cells, leading to an increase of the pH in the stomach. However, no increased risk for the subsequent development of GERD was found so far in patients after H. pylori eradication [25].

1.2.2.

Diagnostics

Invasive and noninvasive techniques are available to diagnose H. pylori infection. The most standardized technique is microscopical examination and cultivation of H. pylori from gastric biopsies (invasive), or a rapid urease test (noninvasive). Molecular tests on biopsies are based on PCR using 16S rRNA or the urease genes as targets. Noninvasive techniques include serology, stool antigen tests or urea breath-tests using non-radioactively labeled urea [16,26].

1.2.3.

Treatment

Eradication of H. pylori infection usually results in cure of the underlying H. pylori-associated disorder [16]. Although H. pylori is susceptible to most antibiotics in vitro [27], many antibiotics are not effective for eradication of H. pylori in infected patients [28]. This failure may be due to the slow growth rate of H. pylori, an inability of antibiotics to attain effective concentrations in the gastric mucus layer [29], or inactivation of antibiotics by the low pH at the site of infection [30]. Therefore, therapy with one antibiotic alone is mostly ineffective. The standard treatment for H. pylori eradication consists of a combination therapy of antibiotics and acid-suppressive drugs. Usually, a triple therapy is given for 7–14 days, consisting of two antibiotics (clarithromycin with either amoxicillin, metronidazole, or tetracycline) together with a proton pump inhibitor (to raise the gastric pH), and/or ranithidine bismuth citrate [16,31]. However, as the prevalence of antibiotic resistance is rising as a consequence of empirical treatment, other strategies, such as vaccination or treatment with inhibitors against essential colonization factors such as bacterial urease, may play a role in the future (see Section 6).

1.3. Mechanisms of Pathogenesis The mucus layer overlaying gastric epithelial cells is the primary niche of H. pylori. This environment is characterized by temporarily acidic pH, active host immune responses and rapid changes in nutrient availability. Therefore, H. pylori requires several systems for colonization and long-term survival in the gastric mucosa. To escape the acidity of the gastric lumen after primary infection, H. pylori moves rapidly to the mucus layer, which is thought to be less acidic (see Met. Ions Life Sci. 2, 545–580 (2007)

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also Section 1.4). H. pylori uses its flagella to reach the mucosa and orients itself with the help of a potent chemotaxis system [32], which, amongst others, responds to the urea gradient spanning the gastric mucus layer [33]. In addition, both flagella and chemotaxis enable H. pylori to adequately react on resource changes. The amount of substrates for bacterial growth and metabolism is variable and the mucus layer itself is thought to be a rather poor source of nutrients [34]. Histological investigations revealed that H. pylori is often located in close proximity to epithelial cells, probably because of the less acidic pH and the local availability of nutrients. To protect itself from peristaltic movements of the mucus and to gain easier access to nutrients, as well as to allow interaction with the host cells, H. pylori expresses adhesins on its surface. Several proteins, like BabA or SabA mediate attachment to Lewisb and Lewisx antigens on epithelial cells [35–37]. Lewisx is an epitope of glycoproteins that is only present in low concentrations in healthy epithelial cells. Upon H. pylori infection, expression of Lewisx antigen is stimulated, leading to increased concentrations of glycoproteins that can be exploited for further H. pylori adherence [35]. It was shown that H. pylori can take advantage of the nutritional exudates leaking from the damaged cells that it adheres to [38]. Thus adhesion is thought to trigger the immune response, leading to enhanced epithelial damage, which finally increases the risk for ulceration, atrophy and gastric cancer [39,40]. H. pylori can actively modulate immune responses predominantly by two major factors that interact directly with host cells: the vacuolating cytotoxin VacA and the cytotoxin-associated protein CagA encoded by the cag pathogenicity island (cag PAI). VacA causes formation and accumulation of endosomal vacuoles in epithelial cells. Furthermore, it forms pores in the cytoplasmic membrane that lead to leakage of anions and urea out of cells. The structure of the vacA gene is highly variable, and certain variants of the VacA protein are associated with increased risk of ulcer disease and cancer [20,21,41–44]. Recently it has been shown that VacA also directly influences T lymphocytes [45]. The genes of the cag PAI code for a type IV secretion system, which injects the CagA protein into host epithelial cells [46]. Inside the cells, CagA is phosphorylated and influences proliferation, apoptosis, cytokine release and cytoskeleton rearrangements. In some gastric epithelial cell lines these alterations lead to a ‘hummingbird’ morphology [20,21,43,44]. Although the biological consequences of the cellular CagA effects are not understood so far, one can speculate that most of them are of advantage for the bacterium. In addition, H. pylori induces inflammatory responses, such as interleukin-8 synthesis [47], and the infection results in a strong accumulation of neutrophils and macrophages in the inflamed gastric mucosa. These innate immune cells play an important role in the defense against bacterial infections, as they are specialized on the eradication of pathogens by phagocytosis. H. pylori is able to interact directly with phagocytes, as it downregulates phagosome formation and persists within macrophages for extended time periods [48–51]. The H. pylori urease, Met. Ions Life Sci. 2, 545–580 (2007)


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which produces carbon dioxide and ammonia from urea (see also Section 2.1), is suspected to play a key role in the interaction with immune cells. Macrophages produce both nitric oxide – created by the inducible nitrite oxide synthase (iNOS)– and superoxides, which in turn react to peroxynitrite [52]. Peroxynitrite can be scavenged and detoxified by carbonic acid, which is an aqueous product of the H. pylori urease reaction [53]. It has been demonstrated that urease induces iNOS [54], which is associated with epithelial cell damage [55] and apoptosis [56]. Furthermore, it is speculated that urease may directly induce cytokine-induced apoptosis of epithelial cells, as the apoptosis levels observed were correlated with H. pylori urease activity. Finally, ammonia released by the urease reaction is able to accelerate apoptosis via tumor necrosis factor α (TNFα) [57,58]. Taken together, these mechanisms indicate that H. pylori is well adapted to host cell interactions and gains access to nutrients by active induction of apoptosis and cell death. Neutrophils and macrophages not only interact directly with bacteria, they are also able to create reactive oxygen species (ROS), like superoxide and hydrogen peroxide. Furthermore, superoxide and hydrogen peroxide are also byproducts of respiration, electron transport, and metal-catalyzed oxidation of metabolites [59]. The dangerous side of superoxides and hydrogen peroxides, which themselves are not very reactive, is that they can form highly reactive hydroxyl radicals. These in turn damage or destroy macromolecules like DNA, proteins or components of the cell wall [60,61]. Hydroxyl radicals can be produced via the Haber–Weiss–Fenton reactions (Equations 1–3): O⫺2 ⫹ Fe3+ → Fe 2⫹ ⫹ O2

(1)

Fe 2⫹ ⫹ H 2 O2 → Fe3⫹ ⫹ OH⫺ ⫹ HO•

(2)

O⫺2 ⫹ H 2 O2 → HO• ⫹ OH⫺ ⫹ O2

(3)

To survive increased ROS production at the site of inflammation, H. pylori expresses several oxidative stress defense mechanisms such as the iron-cofactored superoxide dismutase SodB (HP0389), which detoxifies superoxide radicals into hydrogen peroxide molecules (Equation 4); these are in turn catalyzed to water and molecular oxygen by the catalase KatA (HP0875) (Equation 5) [16,61]. 2 O⫺2 ⫹ 2 H⫹ → H 2 O2 ⫹ O2

(4)

H 2 O2 ⫹ O2 → 2 H 2 O ⫹ O2

(5)

In view of its microaerophilic requirements and low oxygen tolerance, it is not surprising that enzymes involved in the detoxification of ROS are important H. pylori colonization factors. In H. pylori, this is demonstrated by the inability of mutants in genes encoding components or regulators of detoxification enzymes to colonize the gastric environment in animal models [62–68]. Met. Ions Life Sci. 2, 545–580 (2007)

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1.4. The Roles of Urease and Nickel in Gastric Adaptation The acidity in the gastric mucosa is thought to vary around pH 5, with occasional acid shocks occurring when the mucus layer is damaged [69]. This is still under debate, as the proton protective ability of the mucus layer has been questioned [70]. However, despite its acidic habitat, H. pylori is a neutralophilic bacterium, which maintains its cytoplasmic proton concentration at pH 7 [71]. Under in vitro conditions in the absence of urea, the lowest pH allowing growth of H. pylori is about pH 5 [72,73]. H. pylori is able to respond actively to changes in the environmental proton concentrations. At low pH, acidity induces changes in the composition of lipopolysaccharides [74], elevates the expression of chaperone-like proteins [75], and affects transcription of many genes [72,73,76,77]. One of the most important factors to survive the acidic pH in the stomach is the H. pylori urease, which produces ammonia from urea leading to neutralization of gastric acid (see also Figures 2–5). In times of urea shortage H. pylori also possesses alternative systems for ammonia production, like the amidase (HP0294) and formamidase (HP1238) enzymes [72,73]. The fact that urease is a nickel-containing enzyme indicates that maintaining proper nickel metabolism is absolutely essential for the survival in the gastric environment. It was previously demonstrated that transcriptional regulation of the urease structural genes, as well as induction of urease activity, is regulated by the nickel responsive regulator NikR (see also Figures 4–5) and depends on the nickel cofactor concentration, as outlined in more detail below [72,73,78,79]. The importance of urease and nickel metabolism for H. pylori is demonstrated by the inability of urease mutants to colonize the gastric mucosa of mice, gnotobiotic piglets or cynomolgus monkeys [80–85]. And even when the pH of the stomach of gnotobiotic piglets was neutralized by use of proton pump inhibitors, only very low numbers of urease mutants were recovered, indicating that acid neutralization is not the only function of the urease enzyme [84].

1.5. Mechanisms of Metal Ion Homeostasis Metal ions, like iron and nickel, favor the creation of ROS, but are also essential for urease-mediated acid resistance and for bacterial metabolism. Therefore, detoxification of ROS by H. pylori needs to be tightly coupled to the control of intracellular metal ion concentrations, which secures metal ion homeostasis by regulation of metal ion uptake, export and/or storage. While most metal ions are small enough to enter the periplasm via unspecific pores [86], transport over the ion-tight cytoplasma membrane is mediated by specific uptake systems. In bacteria, metal homeostasis is mostly realized by transcriptional regulation via ion-specific regulatory proteins, which combine detector and effector functions in Met. Ions Life Sci. 2, 545–580 (2007)


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one molecule by sensing the intracellular metal concentration and modulation of transcription by DNA binding. When activated, the regulator induces or represses transcription of the corresponding ion uptake, ion efflux and/or ion storage genes. In H. pylori, only two metal regulatory proteins have been identified to date, the nickel-responsive regulator NikR [79] (described in more detail in Section 4.1) and the iron-uptake regulator Fur [87]. The iron-responsive regulatory protein Fur has been found so far in all gramnegative and many gram-positive bacteria, as well as in cyanobacteria [88]. It was first identified in E. coli mutants, which overexpressed siderophores [89]. Therefore, the classical function of Fur is iron uptake regulation. Fur possesses a helix–turn–helix N-terminal DNA binding domain and a C-terminal oligomerization and metal binding domain [90]. Native Fur forms a homodimer composed of 17 kDa subunits. To repress transcription of its target genes, Fur binds ferrous ions as cofactor [91]. Upon binding of Fe2⫹ Fur changes its conformation, which then has an increased affinity for its DNA binding site in the promoter regions of target genes, called Fur boxes [92]. Fur boxes are mostly located around the ⫺10 and ⫺35 region of Fur-repressed genes. When the concentration of ferrous ions in the cell decreases, Fur loses its cofactor and subsequently leaves the Fur box [91]. The E. coli Fur box was first identified as a 19 bp palindromic region (GATAATGAT(a/t)ATCATTATC), but has been reinterpreted as a region of multiple repeats of NAT(a/t)AT [91]. Apart from the classical repressor function, Fur has also been described in E. coli as an activator of gene transcription. Several genes, such as the ferritins bfr and ftn, the aconitase acnA, or the superoxide dismutase sodB, are Fur- and iron-dependent upregulated. Interestingly, these genes do not possess an apparent Fur box [89,91]. However, regulation of these genes is indirect. Fur represses the transcription of the small RNA ryhB, which acts as a Fe2⫹–Fur-repressed negative transcriptional regulator [93]. In the absence of Fe2⫹, Fur is inactive and the small RNA ryhB is transcribed, thus repressing transcription of its target genes. Upon increasing ferrous ion concentrations in the cell, a Fe2⫹–Fur complex is formed, repressing transcription of ryhB and thereby inducing transcription of ryhB target genes [93]. H. pylori also possesses a Fur protein [8,9], which was demonstrated to partially complement a fur mutant of E. coli [87,94], suggesting that H. pylori Fur and E. coli Fur both behave similarly. Interestingly, comparison between the sequences of H. pylori Fur and Fur of Pseudomonas aeroginosa [90] revealed a great homology in the oligomerization domain, but only a low homology in the DNA-binding domain [92], suggesting that the consensus H. pylori Fur box differs significantly from that of other bacteria. H. pylori Fur directly regulates sets of target genes in two different manners. The classic iron-dependent regulation was confirmed in several studies where Fur binds Fe2⫹ and represses the iron uptake by binding to Fur boxes on its iron uptake genes [94–97]. The second manner of direct regulation by Fur was demonstrated with the Met. Ions Life Sci. 2, 545–580 (2007)

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H. pylori pfr gene, which encodes the iron storage protein ferritin (pfr, HP0653) [98] and the sodB gene, encoding an iron-cofactored superoxide dismutase [99]. With the help of DNaseI footprinting experiments, it was demonstrated that pfr [97] and sodB [100] are directly repressed by the iron-free form of Fur. Apart from regulating metal uptake and storage, Fur of H. pylori has also been implicated to be involved in regulation of acid resistance [72,73,101], nitrogen metabolism [102], and oxidative stress resistance [63,64]. In H. pylori, Fur regulates its own gene, and Fur functions are linked to nickel metabolism by the fact that the fur gene is regulated by the nickel responsive regulator NikR [72,73,92,103,104]. Additionally, H. pylori iron uptake [105], regulation [73], and storage [106] are necessary for colonization, as demonstrated in Mongolian gerbil-based and mouse animal models.

2. 2.1.

NICKEL ENZYMES AND ENVIRONMENTAL ADAPTATION Urease

Nickel-containing ureases (urea amidohydrolase, EC 3.5.1.5) are present in bacteria, plants and animals [107–111]. Ureases catalyze urea hydrolysis into ammonia and carbamate (Equation 6), which decomposes spontaneously into another molecule of ammonia and carbon dioxide (Equation 7). H 2 N-CO-NH 2 ⫹ H 2 O → NH 3 ⫹ H 2 N-COOH

(6)

H 2 N-COOH → NH 3 ⫹ CO2

(7)

Carbon dioxide is subsequently converted into carbonic acid mediated by the periplasmic and cytosolic carbonic anhydrases Cah (HP1186) [70] and IcfA (HP0004). Both carbonic anhydrases were demonstrated to be required for urease activity [70,112]. CO2 ⫹ H 2 O → H 2 CO3

(8)

In aqueous solutions, protonation of ammonia and carbonic acid in equilibrium occurs: 2 NH 3 ⫹ 2 H⫹ 2 NH⫹4

(9)

H 2 CO3 H⫹ ⫹ HCO⫺3

(10)

The protonation of ammonia leads to an increase in pH, and the NH3/ NH⫹4 couple reaches its equilibrium at pH 9.2. The H2CO3/HCO⫺3 couple is in Met. Ions Life Sci. 2, 545–580 (2007)


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equilibrium at pH 6.1 [70]. In most organisms, including H. pylori [113], urease activity functions to produce ammonia for nitrogen metabolism. However, as the protonation of ammonia leads to an increase in pH, the urease activity in H. pylori is primarily used to neutralize the acidic surrounding of the human stomach. Activity of the H. pylori urease enzyme does not differ essentially from ureases of other bacteria [109,110]. However, as up to 10% of the whole cell protein can consist of urease, the enzyme activity in H. pylori is 10–100-fold higher than in most other urease-positive bacteria [72,114]. A Km of H. pylori urease between 0.17 to 0.48 mM (urea/min/mg protein) indicates that under physiological conditions H. pylori urease is saturated, as the physiological urea concentration in the gastric mucosa is around 1.7–3.4 mM [110]. In comparison, the Km of ureases of microorganisms, which are exposed to high concentrations of urea, is usually higher [109]. For example, Km values between 13 and 130 mM have been observed in Proteus mirabilis, Sporosarcina ureae and Bacillus pasteurii [109]. Urease of H. pylori is composed of two subunits, UreA (27 kDa) and UreB (62 kDa) [110,115,116]. In the quaternary structure, the subunits form a multimeric enzyme complex with spherical assembly. The native urease holoenzyme is a dodecameric protein structure, consisting of four UreAB trimers ((AB)3) 4 [115,116]. Each UreAB subunit needs two nickel ions for activity [117]. Therefore, 24 nickel ions are necessary for a native ((UreAB)3) 4 holoenzyme. The urease genes are evolutionary related and share a common ancestor. In general, ureases consist of the structural subunits UreABC [109]. In H. pylori, the ancestral ureA and ureB genes are fused and create ureA, whereas the ancestral ureC gene is unfortunately labeled ureB in this bacterium (Figure 2). The H. pylori ureA and ureB (hp0073-hp0072) structural genes for urease subunits are located in a large gene cluster, together with the ureIEFGH genes (Figure 2). The ureABIEFGH operon is coordinately expressed from two different promoters situated in front of the ureA and ureI genes [118]. The ureEFGH genes (hp0070-hp0069-hp0068-hp0067) code for urease accessory proteins, which mediate proper formation of the complex quaternary structure and transport nickel ions into the urease active center.

Figure 2. Genetic organization of the urease operon in H. pylori and Bacillus sp. TB90. The ureAB genes of the ancestral urease operon are fused and labeled ureA, the ancestral ureC gene is labeled ureB in H. pylori. Met. Ions Life Sci. 2, 545–580 (2007)

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The ureI gene codes for a pH-regulated urea channel situated in the cytoplasmic membrane (see Figures 3 and 4). When the periplasmic pH drops below pH 6.1, protons bind to protonable amino acid residues in the periplasmic part of the UreI transporter, presumably leading to conformational changes in the transmembrane domain [119,120]. The UreI channel opens and environmental urea molecules enter the cell [121]. Preliminary experiments indicate that UreI and UreA might interact, suggesting that urea hydrolysis may take place in close proximity to the cell wall to allow fast diffusion of ammonia and CO2 into the periplasm to effectively protect H. pylori against the acidic environment [122]. UreI was thought to be unique for Helicobacter species [110], however, recent publications described UreI urea transporters in Streptococcus thermophilus and Streptococcus salivarius [124]. The accessory proteins UreEFGH incorporate the nickel cofactor into apo-urease and it was shown that urease activity was lost or diminished in accessory gene mutants [109]. The UreH protein of H. pylori, which is homologous to UreD in other bacteria [109], was proposed to be a chaperone to facilitate proper assembly of the urease metallocenter, as interaction between UreD and the apourease in the presence of CO2 was elucidated in Klebsiella aerogenes [125,126]. In H. pylori, interaction was described between UreH and UreF [122]. The function of UreF is still unknown. The accessory proteins HypAB, normally involved in incorporation of nickel into the H. pylori hydrogenase, were also shown to be necessary for the activation of urease [127–129]. The HypB protein is a homolog of UreG, an enzyme that possesses GTP-hydrolyzing activity [128–130]. Furthermore, interaction between UreG and UreE was demonstrated [122,131]. H. pylori UreE is a metallochaperone, which sequesters nickel, although it lacks the histidine rich C-terminal domain present in other UreE proteins [132]. The H. pylori GroES homolog HspA (HP0011) was also suggested to be involved in urease regulation, as it contains a histidine- and glutamine-rich C-terminus [133,134]. An E. coli strain, harboring plasmids with the urease gene cluster and the hspA gene, displayed significantly higher urease activity under low nickel concentrations compared with absence of nickel [133,134], and therefore it was concluded that HspA is a carrier for nickel incorporation into urease [133,134]. It is now thought that the major part of urease is localized in the H. pylori cytoplasm and is released to the environment upon cell death and lysis. In aging cultures, urease was detected on the surface of viable H. pylori cells [135,136]. The relevance of this finding is being debated, as it may represent an artefact of cell lysis in vitro [137].

2.2. The Role of Urease in Acid Resistance Urease converts urea into ammonia and carbon dioxide (Equations 6 and 7), which can both be protonated (Equations 8 and 9). With an equilibrium at pH 9.2, the buffering capacity of the NH3/NH⫹4 couple is too small to maintain Met. Ions Life Sci. 2, 545–580 (2007)


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Figure 3. Model of acid resistance in H. pylori (modified from [71]). Urea enters the periplasm via pores in the outer membrane (OM). The inner membrane (IM) transporter UreI transports urea under acidic conditions into the cytoplasm. Here, urea is converted into NH3 and CO2 by the urease, and the gases diffuse back into the periplasm. There, the α-carbonic anhydrase hydrates CO2 to H2CO3, which dissociates into H⫹ and HCO3⫺. The created proton is accepted by one NH3 to create NH⫹4 , leaving the second NH3 to neutralize a proton from acid leaking into the periplasm.

the cytoplasm at neutral pH [70]. Because the H2CO3/HCO⫺3 equilibrium is with pH of 6.1 closer to cytoplasmic pH, the following model for interactions between urease and carbonic anhydrase in acid resistance was proposed (Figure 3): Urea is hydrolyzed in the cytoplasm into two molecules of ammonia and one molecule of carbon dioxide. All products diffuse into the periplasm, where the carbonic anhydrase converts CO2 into H2CO3, which dissociates into HCO⫺3 and H⫹ (Fig. 3). The proton is detoxified by one molecule of ammonia, whereas the second molecule of ammonia can be used to neutralize protons entering the periplasm from the surrounding medium [70]. Transcriptional regulation of bacterial ureases is usually mediated by nitrogen, urea, growth phase or pH [107,110]. In H. pylori, urease gene transcription is not altered upon changes in nitrogen and urea level. Therefore, it was thought for a long time that urease expression was constitutive and thus not regulated at all. However, only recently, it was demonstrated that transcription of the H. pylori urease is acid-regulated [72,73,76,118,138]. Northern hybridization analysis revealed elevated levels of mRNA fragments that contain the ureAB and ureIE’ genes [72,73,118], respectively, in cells incubated for 30 min in medium at pH 6 [72,73,118]. Urease transcription was furthermore shown to be nickel-induced via the nickel regulator NikR (Table 1; see Section 4) [72,78,79,139,140]. This regulation makes biological sense, as urease activity is limited by the availability of free Met. Ions Life Sci. 2, 545–580 (2007)

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Table 1.

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Mechanisms of urease regulation.

Level

Mechanism

Reference

Transcriptional

Acid-dependent induction Nickel-dependent induction Modification of urease transcripts

[72,73,78,79] [118]

Post-transcriptional

Stabilization of urease mRNA

[118]

Post-translational

Nickel-dependent activation of urease apo-enzyme Acid- and UreI-dependent potentiation of urease activity

[78] [121]

nickel in the cell, and it is believed that the gastric nickel concentration is coupled to the level of acidity (see Section 4) [72,104]. Recently, the two-component system ArsRS was implicated to regulate urease transcription under acidic conditions. With the help of primer extention experiments, it was shown that both the ureA and ureI promoter are acid- (pH 5) and ArsS-dependent induced [76,138]. Furthermore, it was shown with DNaseI footprint analysis that phosphorylated ArsR can bind to both the ureA and ureI promoters [76]. In H. pylori, urease activity is regulated both by post-transcriptional and posttranslational mechanisms (Table 1). These include the acid-dependent access to the urea substrate due to the opening and closing of the UreI channel, a posttranscriptional stabilization of the urease mRNA under acidic pH and a nickelinduced activation of apo-urease. Ammonia production by urease is relatively low when measured at neutral pH. However, there is a 10–20-fold induction of ammonia production by urease when the pH is decreased to 5.5 or lower. This induction of ammonia production was demonstrated to be due to increased influx of urea, as mediated by the pH-dependent opening of the UreI channel, which has increased urea permeability at low pH. The UreI channel opens at pH 6.1, and activity was maintained down to a pH as low as pH 2.5 [137,141]. At neutral pH, UreI is closed to circumvent the raise of the cytoplasmic pH above neutral [121]. A second post-transcriptional induction mechanism of urease was reported to be pH-dependent mRNA stability at low pH, as urease mRNA is stabilized at acidic pH. Additionally, under acidic conditions, higher levels of mRNA transcripts were found, as well as different transcripts, especially of the mRNA of the urease accessory genes [118]. It was concluded that the posttranscriptional stabilization of the accessory genes increased the efficiency of the urease assembly [118]. The third mechanism is post-transcriptional induction of urease activity at the protein level. After nickel-supplementation of growth medium the increase in urease activity was demonstrated to be much higher than the raise in protein Met. Ions Life Sci. 2, 545–580 (2007)


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levels [78]. Furthermore, a nickel-dependent augmentation of urease activity is still observed when nickel-dependent transcription is blocked [79]. It was proposed that H. pylori can use the nickel concentration as an indicator for low pH, because the bioavailability of nickel ions is higher at low pH [72,78,79,142].

2.3.

Ni/Fe-Hydrogenase

The membrane-bound hydrogenase enzyme (cytochrome c3 oxidoreductase, EC 1.18.99.1) catalyzes the ‘splitting’ of electrons from hydrogen (Equation 11), which are in turn used for energy generating purposes [143]. H2 2 H⫹ ⫹ 2 e⫺

(11)

H. pylori expresses a Ni/Fe-hydrogenase which displays many features common to bacterial Ni/Fe-hydrogenases, using O2 as terminal end electron acceptor [144]. The H. pylori hydrogenase operon consists of five genes: hydA (hp0631) and hydB (hp0632) encode the small 26 kD and the large 65 kD subunit, respectively. The hydC (hp0633) gene codes for a cytochrome c subunit, and the HydD sequence (hp0634) shares similarities with a protease involved in the maturation of the large hydrogenase subunit of Wolinella succinogenes. The function of the hydE (hp0635) gene product is currently unknown [145]. Similar to urease, hydrogenase enzyme activation requires besides nickel and iron cofactors, the HypABCDEF accessory proteins [8,9]. The corresponding genes hypA (hp0869), hypBCD (hp0900-hp0899-hp0898) and hypEF (hp0047hp0048) are located at different places in the chromosome. HypA was shown to have nickel-binding activity, whereas HypB displayed GTPase activity [127,128]. Inactivation of all the hydrogenase accessory genes leads to reduced activity or complete absence of hydrogenase activity [127], and it was demonstrated that the HypA and HypB proteins are also necessary for urease activity [127]. Nickel supplementation of growth medium restored urease activity, but not the hydrogenase activity, of a hypAB mutant [127].

2.4.

Hydrogenase and Gastric Adaptation

The hydrogenase system with O2 as terminal end electron acceptor is proposed to help H. pylori to adapt to the gastric mucosa. This assumption is based on the observation that H. pylori seems to be limited in its use of oxidizable organic compounds, and that the mucus layer of the human stomach is thought to be relatively poor in nutrients [34]. Furthermore, the nature of primary carbon and nutrient sources of H. pylori in the gastric mucosa of the human stomach is still unknown [34,146,147]. Met. Ions Life Sci. 2, 545–580 (2007)

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Hydrogen is produced by colonic bacteria and from there transported via the bloodstream to the gastric mucus layer. H. pylori can efficiently use its hydrogenase system as its Km is 1.8 µ M [34], compared with a hydrogen concentration of 43 µ M in the mucus layer of the stomach of living mice. H. pylori hydrogenase activity is two- to four-fold induced by addition of 10% H2 [34], and as human cells cannot use hydrogen as energy source, there is no competition between H. pylori and its host for hydrogen. Evidence for the use of hydrogenase to adapt to the human stomach was further provided by the fact that H. pylori hydrogenase mutants are deficient in gastric colonization of a mouse model [34].

3.

NICKEL UPTAKE SYSTEMS

The important roles of nickel ions in urease activity, acid resistance, and gastric colonization necessitate the presence of a high-affinity nickel uptake system, which secures nickel availability even at low environmental nickel concentrations. Two classes of bacterial nickel transporters have been described. In E. coli nickel is imported into the cytoplasm via an ABC-transporter, consisting of the NikA periplasmic-binding protein, the inner membrane permeases NikBC and the NikDE ATPase proteins [148]. The E. coli NikABCDE system is transcriptionally regulated by NikR, which is encoded downstream of the nik gene cluster [149]. Orthologs of the Nik ABC-transporter are found in Brucella [150] and Yersinia species [151], and in Actinobacillus pleuropneumoniae [152]. A second class of nickel transporters is represented by the Alcaligenes eutrophus HoxN protein, which is a high-affinity nickel permease [153,154] with eight transmembrane segments [155,156]. Other homologs of HoxN were identified in Bradyrhizobium japonicum [157], Rhodococcus rhodochrous [158], and in thermophilic Bacillus species [123].

3.1.

Nickel Import by the NixA Protein

H. pylori possesses a HoxN-type, nickel-specific uptake system designated NixA (HP1077), which was discovered by the finding that additional nickel was needed to activate recombinant H. pylori urease in an E. coli strain [159]. The nixA gene encodes a monomeric, 37 kDa protein [159], also with eight transmembrane domains, like HoxN [160]. Both the carboxy and amino terminal ends of the protein are situated in the cytoplasm [160], and motives in transmembrane domain II (GX2HAXDADH) and III (GX2FX2GHSSVV) are essential for nickel transport [160,161]. The nickel concentration in the gastric mucosa is presumably similar to the concentration of nickel in the serum, which is 2–11 nM [161,162]. The affinity of NixA for nickel is 11.3 nM, and thus in Met. Ions Life Sci. 2, 545–580 (2007)


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the gastric mucus layer nickel uptake via NixA will be efficient [161], especially as nutrients like tea, chocolate, cornflakes or nuts are rich in nickel [162]. High-affinity nickel uptake is an energy-dependent process, as demonstrated by experiments with the Rhodococcus rhodochrous NixA homologue NhlF. However, in H. pylori the energy source driving nickel import is currently unknown [163,164]. Nickel uptake by NixA is thought to be the major nickel uptake route in H. pylori. Inactivation of nixA resulted in a 75% nickel influx reduction [165]. Competition experiments indicated that other divalent ions such as Co2⫹, Cu2⫹ and Zn2⫹ ions may be cotransported by NixA [161]. The importance of NixA for the colonization of the stomach was demonstrated in vivo. In mice, the gastric colonization capacity of a H. pylori nixA mutant was decreased compared with the wildtype strain, and nixA mutants where outcompeted by the wild-type strain [80].

3.2.

Possible Alternative Nickel Transporters

Two additional putative nickel uptake transporters have been proposed in H. pylori. The first one is the magnesium transporter CorA (HP1344), a single-component cytoplasmic membrane protein. Results from growth experiments suggested that CorA transports Ni2⫹ ions at high environmental nickel concentrations [166]. The nickel transport function of CorA was supported by the finding that an E. coli corA mutant displayed nickel resistance [167], which was abolished by transfer of a plasmid carrying the intact H. pylori corA gene. However, subsequent Mg2⫹ supplementation restored nickel resistance to the E. coli corA mutant carrying H. pylori corA, indicating that Mg2⫹ is the dominant substrate for H. pylori CorA. This was further supported by analysis of H. pylori corA mutants, which displayed no alterations in urease activity or expression, when compared with the wild-type strain [166]. The abcABCD operon may encode for a second alternative nickel transporter. The AbcC protein displays low level of homology to the ATP-binding protein NikD from E. coli, however the other three proteins displayed no significant similarity to any known proteins [168]. The AbcABCD proteins may be involved in nickel uptake, as it was demonstrated that an abcD–nixA double mutant nearly abolished urease activity without altering urease synthesis [165,168]. However, nickel transport experiments with abcD and abcC–nixA double mutants revealed no significant difference in nickel uptake [168]. Recently the dipeptide permease operon (dpp, hp0299–hp0302), which shares similarity with the nik gene cluster of E. coli (DppA, DppC and DppD share respectively 40%, 53% and 56% similarity with the E. coli gene products) was implicated to contribute to urease activity. However, in this publication, it was demonstrated that the dpp operon is not included in urease regulation, as a dppA mutant showed no difference in urease activity [169]. Met. Ions Life Sci. 2, 545–580 (2007)

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MECHANISMS OF NICKEL REGULATION

4.1.

The NikR Regulator

So far, besides H. pylori NikR, the only nickel regulator described and biochemically characterized is E. coli NikR [149,170,171]. However, NikR orthologs have been identified in other gram-negative bacteria and Archaea [79,170,172,173]. NikR is a member of the Arc/CopG/MetJ/Mnt family of ribbon–helix–helix (RHH) family of DNA-binding proteins, which function as transcriptional regulators. E. coli NikR possesses an N-terminal DNA-binding domain, which recognizes a 28 bp palindromic region (GTATGA-N16-TCATAC) in the nikA promoter. The C-terminal domain contains a nickel-binding motif. Binding of nickel ions induces changes in the NikR secondary structure, and thereby enhances DNA binding [170,171,174–176]. Interestingly, E. coli NikR was only shown to regulate nickel uptake by nikA, but other targets of E. coli NikR have not been identified. The H. pylori genome sequence [8,9] allowed identification of a gene for a NikR (HP1338) ortholog, which displays 30% identity and 68% similarity to E. coli NikR [79]. The 17 kDa protein was proposed to act as transcriptional repressor, which senses the cytoplasmic nickel concentration by binding of intracellular free nickel [104]. With the help of gel filtration size exclusion chromatography with recombinant NikR it was demonstrated that, similar to E. coli NikR, H. pylori NikR is of tetrameric structure, independent of the nickel-bound status of the protein [140,174]. Detailed analysis revealed that H. pylori NikR is required for maintaining nickel and iron homeostasis, as well as for regulation of motility and chaperones [72,73,78,79,172]. Furthermore, we have hypothezised that H. pylori NikR functions as master regulator of acid adaptation [104] and the importance of NikR in colonization and gastric adaptation was underlined by the fact that NikR mutants were attenuated in colonizing the mouse stomach [73].

4.2.

Genes Regulated by NikR

In H. pylori, several genes and metabolic pathways have been demonstrated to be regulated by NikR and/or by nickel (Figure 4).

4.2.1.

Regulation of Nickel Uptake

Similar to E. coli NikR, the H. pylori NikR protein controls nickel uptake (Figure 4) via direct nickel-dependent binding to the promoter region of the high-affinity nickel importer gene nixA [139]. However, there is no consensus on the exact mechanism of how NikR exerts its regulatory function. Using array and spot blot analysis, it was suggested that nixA transcription is nickel- and NikR-induced [172]. Met. Ions Life Sci. 2, 545–580 (2007)


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Figure 4. Regulation of nickel metabolism and acid resistance by NikR. Nickel uptake by NixA is nickel-dependently repressed, whereas the urease activity is nickeldependently induced by the nickel-responsive regulator NikR. Further genes regulated by NikR are the fur regulatory gene and possibly arsRS and flbA. Increased nickel availability is thought to be a signal for a low pH of the environment. Transcription of urease is induced to convert urea which is entering through UreI under low pH into NH3 and CO2. OM: outer membrane; IM: inner membrane.

However, nickel-induced transcription of nixA would result in continuous influx of nickel in wild-type cells, and this is contradicted by the high-level nickel resistance of the wild-type strain and by the nickel sensitivity of a nikR mutant [79,172]. Furthermore, mutation of the nixA gene was shown to complement nickel sensitivity of an H. pylori nikR mutant [139]. With the help of DNaseI footprint analysis using recombinant NikR nickel- and NikR-dependent repression of the nixA gene by binding to an operator sequence overlapping the ⫺10 and ⫹1 sequence was demonstrated [139]. Interestingly, the binding sequence recognized by H. pylori NikR in the nixA promoter differs significantly from the binding sequence recognized by E. coli NikR.

4.2.2.

Control of Urease Synthesis

Another promoter directly regulated by H. pylori NikR is the ureA promoter. Urease expression is induced by nickel at the transcriptional level (Figure 4), and Met. Ions Life Sci. 2, 545–580 (2007)

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mutational analysis revealed that NikR is involved in the transcription of urease genes [72,73,78,79,140]. Gelshift and DNaseI footprint experiments allowed the identification of the NikR target sequence at approximately nucleotides ⫺50 to ⫺90 upstream of the canonical σ80 site in the ureA promoter [139,140]. The binding sequence displayed a partial overlap with an imperfect palindromic region at position ⫺49 to ⫺69 upstream of the transcriptional start site of the ureA gene [139,140]. It was hypothesized that NikR binding to this region would give easier access for the RNA polymerase to bind to the promoter [79], however the exact role of this palindrome is still not clear. With the help of scanning mutagenesis of the ureA promoter a 26 bp operator sequence was found intersected with 9 nucleotides not necessary for binding [140]. As the NikR binding sites in the ureA and nixA and other genes revealed no similarities besides an AT-rich sequence [139,140], further analysis is necessary to define a NikR box in H. pylori.

4.2.3.

Modulation of Fur and Amidase Genes

NikR forms part of a complex regulatory network, as it modulates iron metabolism and ammonia production by amidases in response to acid and nickel (Figure 5). In general, amidases catalyze the conversion of amide substrates to the corresponding carboxylic acid, and ammonia, however both amidases use different substrates. AmiE hydrolyses propionamide, acetamide, and acrylamide, but

Figure 5. Transcription of ammonia-producing genes is mediated via a regulatory cascade involving Fur and NikR. Transcription of urease is induced, whereas transcription of the iron-uptake regulator Fur is repressed by NikR. The Fur regulator in turn represses transcription of the amidase gene, while regulation of the formamidase gene by Fur involves a currently unknown mediator. Met. Ions Life Sci. 2, 545–580 (2007)


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not formamide, whereas AmiF exclusively hydrolyses formamide [177,178]. It is thought that nickel bioavailability may be an indicator for sensing the environmental pH [72]. Therefore, it has been demonstrated that under mild acidic conditions control of both the amidase and the formamidase genes is mediated in a regulatory cascade by NikR and a second metal regulatory protein, the iron uptake regulator Fur [73,104]. NikR represses the fur gene (Fig. 5), whose gene product, the iron-uptake regulator Fur [73,104,172] directly represses the amiE gene and indirectly, over some unknown mediator, represses the formamidase gene amiF (Fig. 5).

4.2.4.

Regulation of the NikR Gene

The H. pylori NikR protein displays nickel-dependent autoregulation by repressing the promoter of its own gene [140,172]. This is in contrast to the E. coli NikR protein, which is so far only involved in the nickel-dependent regulation of the nikABCDE operon, but not involved in nickel-dependent autoregulation [140,149]. Quite recently it was also demonstrated that H. pylori NikR binds in competition with Fur to the (different, but overlapping) operator sites in the nikR and also in the exbB genes [140]. Additionally, the fact that transcription of the nikR gene is induced by acid provides evidence that a reduced pH leads to a better nickel bioavailability for H. pylori [104].

4.2.5.

Hydrogenase and Other Genes

NikR may regulate H. pylori hydrogenase, as transcription of the hydrogenase operon was repressed in a nikR mutant, but induced in a H. pylori fur mutant, suggesting that transcriptional regulation of hydrogenase is similar to the amidases [72,73,172]. In E. coli, NikABCDE and hydrogenase expression levels are closely linked. Furthermore, when the nickel concentration is high, the nikABCDE operon is under complex regulation involving NikR [148,179]. Other genes that displayed NikR-dependent regulation are involved in respiration, stress response or chemotaxis [172]. These genes can be triggered by the high nickel concentration, or, as nickel creates stress, these genes are induced as part of the general defense system.

4.3.

Metal Metabolism and Acid Regulation

Consistent with its restricted host range and single target organ, the H. pylori genome contains a relatively low number of regulatory proteins. However, three of them are involved in acid regulation: the nickel responsive regulator NikR; the ferric uptake regulator Fur; and the two-component regulatory system Met. Ions Life Sci. 2, 545–580 (2007)

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ArsRS (HP0166/0165) [73,76,104]. Mediation of acid regulation might occur via the cascade NikR⬎Fur⬎ArsR (Figure 4), as expression of Fur and ArsR proteins is acid-repressed [104], and expression of ArsR is absent in a fur mutant background [180]. It is thought that H. pylori is detecting changes in the external pH through sensing the availability of nickel ions in the cytoplasm, as the solubility of metal ions, like nickel and iron, is strongly reinforced under acidic conditions [104,142]. The acid-mediated solubility of metal ions to enter the cell is thought to be increased: (a) due to increased bioavailability of nickel under acidic conditions; (b) increased expressions of nickel transporters; or (c) increased efficiency of the nickel transporter NixA [104]. The following is a model (Figure 4) of acid regulation via the NikR ⬎ Fur ⬎ ArsR cascade [104]. Transcription of urease genes occurs, even when it can not be activated by nickel [139]. Under low nickel concentrations imported free nickel ions will activate the pool of preformed urease apo-enzyme. When under higher nickel ion concentration all urease apo-enzyme is activated, further imported free nickel ions are available in the cytoplasm. This leads to formation of NikR–nickel complexes. These NikR–nickel complexes subsequently bind to target promoters with different affinities. At first, repression of the nixA promoter will occur, even at low nickel concentrations, and therefore lead to decrease and subsequently cessation of NixA-mediated nickel uptake. The upstream binding site for NikR in the ureAB operon is supposed to have a lower binding affinity compared with the nixA promoter. As a consequence, it will be induced when the intracellular nickel concentration further increases, leading to an induction of transcription of the ureAB genes. Moreover, NikR will bind to the promoter sites of other target genes with low-binding-affinity sites in genes for other regulators such as the fur gene and probably also the arsR, hspR, and flbA genes. Recent observations [76] favor an ArsRS-dependent regulation of the acid regulation of H. pylori, as a nikR-mutant in H. pylori G27 did not display acid-dependent regulation of the urease genes as previously described [104]. However, it has yet to be demonstrated whether and how ArsRS senses changes in acidity, as the experiments demonstrating direct acid-dependent regulation of ureA and ureI by ArsRS were performed in vitro using acetylphosphate [76].

5. 5.1.

PROTECTION OF NICKEL METABOLISM Nickel Binding and Storage

As nickel ions are a necessity for H. pylori to survive in the stomach, and because nickel availability may drastically change, nickel storage would be beneficial for H. pylori. The hydrophilic H. pylori nickel-binding protein Hpn (HP1427) could perform corresponding functions. The 180 bp of the hpn gene encode for a 7 kD protein accounting for approximately 2% of the whole cell protein [181]. Nearly Met. Ions Life Sci. 2, 545–580 (2007)


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half (47.6%) of its 60 aa are histidines (28 aa), which have a high affinity for nickel. An hpn mutant was more susceptible for Ni2⫹ added to the medium compared to the wild-type [181,182]. Hpn was shown to exist mainly as 20-mer and is able to bind five nickel ions per monomer. The binding affinity can be reversed, as demonstrated with added EDTA, and the pH for half dissociation of nickel ions is pH 6.3. Binding of nickel ions changed the conformation of the protein. As the nickel ion concentration is higher in a H. pylori 26695 wild-type compared with a hpn mutant, it was concluded, that Hpn might be a storage and detoxification system for excess nickel ions in the cell [183]. Furthermore, transcriptome analysis demonstrated a nickel- and NikR-dependent regulation of hpn [172], as well as an acid-induced induction [73]. Transcription of hpn is also affected by iron in a Fur-dependent fashion [184]. The putative glutamine-rich protein HP1432 (16 out of 72 aa are histidines, clustered in 3 domains) – a homolog of Hpn – might also be involved in nickel binding. The regulation of HP1432 is similar to hpn, as it is nickel- and NikRdependent, as well iron- and Fur-dependent regulated [73,172,184]. In addition, HP1432 is induced at the transcriptional level via ArsRS [138,185].

5.2.

Specific Export of Other Metal Ions

Depending on the diet, the gastric mucosa represents a highly variable natural habitat, in which changes in the environmental metal ion concentration occur within minutes. Alterations in the metal ion availability occur by release from food or by the cation-chelating activity of gastric mucus or host proteins [186]. The finding that H. pylori urease is strongly inhibited by iron, copper, or zinc in vitro [187] indicated that the gastric pathogen requires specific mechanisms, which protect urease and nickel metabolism from interferences with other cations. The average daily requirement for trace metals and the ionic content of drinking water, allows the estimate that H. pylori is exposed to iron, zinc, and copper in the micromolar range. Thus, for continuous persistence in the human stomach, H. pylori has evolved an extended repertoire of adaptive mechanisms to maintain cytoplasmic metal ion homeostasis and urease activity, even if the environmental conditions change drastically. Therefore, H. pylori contains genes for a multitude of metal ion transport systems, which differ in regulation and ion specificity [8,9]. In bacteria metal ions are usually detoxified by efflux via resistance-nodulation-cell division (RND)-type exporters, cation diffusion facilitators (CDF), and P-type ATPases (for review, see [188]). The P-type ATPases and CDF proteins transport ions from the cytoplasm to the periplasmic space. In other bacteria, the proton-driven RND-type metal efflux pump Czc composed of the inner membrane, periplasmic, and outer membrane proteins CzcA, -B and -C, respectively, mediate metal ion export to the environment [189, 190]. It was experimentally proven that H. pylori exports copper, nickel, zinc, cobalt, and cadmium ions from Met. Ions Life Sci. 2, 545–580 (2007)

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the cytoplasm to the periplasmic space via the P-type ATPases CopA [191–193] and CadA [194,195]. In addition, genes for two Czc-type exporters are present in the H. pylori genome [8,9]. The czcB homologs hp0970 and hp1328 are located directly upstream from the corresponding czcA homologs hp0969 and hp1329. Whereas metal export functions of HP0969/HP0970 have not been investigated in detail so far, it was recently demonstrated that the czcAB gene pair hp01328/ hp1329 forms, together with the flanking crdA and crdB genes, an RND-type copper resistance determinant [196]. Transcription of the crdA gene is strongly induced by environmental copper ions and this regulation is mediated by the CrdRS two-component regulator pair [197a]. The essential role of copper regulation in the gastric environment was underlined by the earlier finding that H. pylori mutants deficient in CrdRS did not colonize a mouse infection model [197b].

6. METAL METABOLISM AS DRUG TARGET: THERAPEUTIC CONSIDERATIONS The most effective treatment for H. pylori infection is an antibiotic triple therapy. However, the prevalence of antibiotic resistance is rising, especially in developing countries, where most antibiotics can be obtained without special prescription [198]. Because clarithromycin and metronidazole are commonly used for H. pylori eradication, it is not surprising that in Europe approximately 10% and even 30% of H. pylori isolates display clarithromycin or metronidazole resistance, respectively. In developing countries the numbers of resistant isolates are increased to 25–50% for clarithromycin and to ⬎ 90% for metronidazole [199–201]. Therefore, it is necessary to develop new strategies for treatment of H. pylori infections. As H. pylori uses the nickel-cofactored urease enzyme as first-line defense for acid resistance and gastric colonization, the nickel metabolism represents a potential novel drug target. Three possible new lines of treatment of H. pylori infection will be discussed: (i) vaccines against antigens of the metal metabolism or metal-cofactored proteins such as UreAB [202]; (ii) use of inhibitors against essential enzymes of the H. pylori metal metabolism; and (iii) a metal diet.

6.1. Vaccination One of the best ways to battle pathogens is the use of vaccination. Two possible vaccination strategies can be envisioned for therapy of H. pylori infection: A prophylactic vaccine, where antigens are provided prior to infection in order to try and prevent new infections, and a therapeutic vaccine, which is used to treat existing infections. The most suitable antigens for both types of vaccines are thought to be abundantly expressed and well conserved surface-exposed proteins. The UreAB and HspAB subunits are cytoplasmic proteins; however, both have Met. Ions Life Sci. 2, 545–580 (2007)


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been described to be surface-bound in stationary H. pylori cultures [136]. In addition, urease is an important virulence factor and therefore these proteins have already been used in several vaccination experiments. Prophylactic vaccination with these recombinant proteins induced good protection against H. pylori and H. felis in mouse models, however, they failed or at best gave modest protection in monkeys or humans [202]. Also vaccination with a triple vaccine consisting of recombinant NapA (an iron-binding protein that shows homology to the Dps family of iron-binding proteins [63]), CagA, and VacA only showed moderate protection of beagle dogs [203]. So far, it is unknown why vaccines are quite effective in a rodent model, but not in humans. An explanation might be that H. pylori is able to alter the human immune response against itself in humans, but these mechanisms might not be as effective in its non-natural host [204].

6.2.

Inhibition of Metalloenzymes

The availability of whole genome sequences, transcriptomics and proteomics made it possible to identify genes unique for H. pylori as subsequent growth and colonization experiments with mutant strains identify genes essential to H. pylori. With further analysis of the regulation and mechanism of action, it may be feasible to identify possible targets for inhibitors, which can in turn alter expression of essential genes or block enzyme activity. Possible targets for inhibitors are the H. pylori hydrogenase and urease systems including accessory proteins. Both nickelcofactored proteins are absent in humans, but are present in the H. pylori periplasm (hydrogenase and UreI) or cytoplasm (urease). H. pylori mutants deficient in urease or hydrogenase were unable to colonize the stomach in animal models [110,127] and both enzymes have unique active centers and share similar mechanisms for activation by accessory proteins. Therefore, the design of inhibitors of hydrogenase, urease enzymes or corresponding accessory proteins may represent a promising strategy for the development of novel anti H. pylori agents [109,144,147].

6.3. Metal Diet The use of nickel as a cofactor for the virulence factor urease creates a strong need for H. pylori to acquire environmental nickel ions. Because nickel ions are not used by the human metabolism [205] a nickel-deficient diet might help to treat H. pylori infection. The results from analysis of the nickel metabolism have demonstrated that nickel depletion or probably even a nickel shortage is detrimental for H. pylori growth and for maintaining acid resistance required for effective gastric colonization [78,79]. However, several human food sources such as coffee, tea, nuts, and chocolate are rich in nickel ions [162]. Met. Ions Life Sci. 2, 545–580 (2007)

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CONCLUSIONS

Although H. pylori colonizes a hostile environment, this bacterium is exquisitely well adapted to the conditions in the gastric mucosa. The bacterium can overcome shortage of nutrients by turning immune defense mechanisms against the host itself, leading to damage of epithelial cells and release of nutrients. A second strategy to overcome nutrient shortage used by H. pylori is to use resources, which are not in competition with its host. For example, H. pylori uses hydrogen to gain energy, a resource that humans can not use. Furthermore, it uses urea, a waste-product of the nitrogen metabolism of humans, to survive the acidity of its niche. For both processes, the energy metabolism and the acid adaptation nickel-cofactored enzymes are necessary, again something that is not used by humans. However, due to the fact that nutrients used by humans are rich in nickel, there will be no nickel shortage in the gastric environment. H. pylori has a paucity of regulatory proteins. From the few regulators identified and characterized at present, the nickel responsive regulator NikR seems to be of high priority, as many metabolic pathways essential for gastric adaptation are regulated by NikR. Furthermore, NikR may be a master regulator of the acid metabolism, which is orchestrated by a complex regulator cascade. Furthermore, H. pylori may use the nickel concentration as indicator for acidity, as nickel bioavailability is greatly enhanced at low pH. Finally, as nickel ions are not used in the human metabolism, the nickel metabolism of H. pylori could represent a novel drug target for inhibitors. However, nickel-cofactored urease and hydrogenase enzymes essential for gastric survival are activated and regulated in a complicated way, indicating that further research is necessary to unravel the role of nickel in gastric adaptation or to make use of the susceptible nickel metabolism for the development of novel anti-H. pylori therapies.

ABBREVIATIONS ATP CDF EDTA Fur GERD GTP iNOS NikR ORF PCR RHH

adenosine 5′-triphosphate cation diffucion facilitator ethylenediamine-N,N,N′,N′-tetraacetate iron-uptake regulator gastroesophageal reflux disease guanosine 5′-triphosphate inducible nitrite oxide synthase nickel responsive regulator open reading frame polymerase chain reaction ribbon–helix–helix

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RNS ROS SodB TNFα WHO

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resistance-nodulation-cell division reactive oxygen species superoxide dismutase B tumor necrosis factor α World Health Organization

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16 Nickel-Dependent Gene Expression Konstantin Salnikow* and Kazimierz S. Kasprzak Laboratory of Comparative Carcinogenesis, National Cancer Institute at Frederick, Frederick, MD 21702, USA <[email protected]>

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4. 5. 6.

INTRODUCTION 1.1. Health-Related Effects of Nickel Exposure 1.2. Alteration in Gene Expression as the Molecular Basis of Nickel-Induced Health Effects GENETIC AND EPIGENETIC CHANGES IN NICKEL-EXPOSED CELLS 2.1. DNA Methylation and Transcription 2.2. Histone Modifications and Transcription ALTERATION OF GENE EXPRESSION FOLLOWING NICKEL-INDUCED LUNG INJURY 3.1. Genes Coding for Metal-Binding Proteins 3.2. Surfactant Genes Expression 3.3. Genes Coding for Extracellular Matrix Proteins 3.4. Genes Coding for Cytokines and Chemokines NICKEL-INDUCED ALLERGY AND GENE EXPRESSION NICKEL-INDUCED EXPRESSION OF ERYTHROPOIETIN ALTERATION OF TRANSCRIPTION FACTORS AND SIGNALING PATHWAYS 6.1. Protein Hydroxylation and Ascorbate 6.1.1. HIFα Hydroxylation and Transcriptional Response 6.1.2. Extracellular Matrix and Transcription 6.2. ATF-1



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Nickel-Induced Oxidative Stress 6.3.1. NF-κ B 6.3.2. AP-1 6.4. Nickel-Induced Changes in Calcium Homeostasis 6.4.1. NF-AT 6.5. PI3 Kinase and Downstream Transcription Factors 6.6. DNA-Damage Response and Apoptosis 6.6.1. P53 6.7. Retinoblastoma 6.8. FHIT 7. CHANGES IN GENE EXPRESSION AND NICKEL CARCINOGENESIS 8. CONCLUSIONS ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES

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INTRODUCTION Health-Related Effects of Nickel Exposure

Nickel is widely used in modern industry with other metals to form alloys to produce coins, jewelry, and stainless steel. It is also used for nickel plating, batteries production, and as a catalyst. Lately, it turns out to be a good catalyst for the production of carbon nanotubes. This new technology will further increase consumption of nickel compounds. Growing demand for this metal and its compounds inevitably causes increased production, recycling, and disposal. Workers are exposed to nickel at all stages of the processing of nickel-containing products through air, water or skin contacts. However, inhalation is the main and most important route of exposure. The exposure to airborne nickel-containing particles has long been known to cause acute respiratory symptoms or illness, ranging from mild irritation and inflammation of the respiratory system to bronchitis, pulmonary fibrosis, asthma, and pulmonary edema [1]. Additionally, nickel exposure also may cause cardiovascular and kidney diseases, as well as allergic contact dermatitis. Yet, the most serious concerns are related to nickel carcinogenic activity. These concerns represent an area of considerable research interest and activity. Possible mechanisms of nickel carcinogenesis are also discussed by Kasprzak and Met. Ions Life Sci. 2, 581–618 (2007)

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Salnikow in Chapter 17 of this book and in a number of review papers published in recent years [2–6]. Epidemiological studies have clearly implicated nickel compounds as human carcinogens based upon a higher incidence of lung and nasal cancer among nickel mining, smelting, and refinery workers [7,8]. Welding fumes, which contain significant levels of nickel have been shown to induce more lung injury and inflammation than mild steel welding fumes, which contain mostly iron [9]. The health effects of inhalational exposure to soluble and insoluble nickel compounds are summarized in a number of recent publications [1,3,8–12]. Acute lung injury following nickel exposure was demonstrated in mice and rats [13–16] and in some instances this injury resulted in death due to endothelial disruption and hemorrhagic edema [17]. In various animal models, chronic exposure to nickel compounds induce tumors at virtually any site of administration [2,6]. Nickel compounds are efficient transforming agents for rodent and human cells in vitro [18–21]. Based on these observations the International Agency for Research on Cancer (IARC) evaluated the carcinogenicity of nickel in 1990 [22] and concluded that all nickel compounds except for metallic nickel are human carcinogens.

1.2. Alteration in Gene Expression as the Molecular Basis of Nickel-Induced Health Effects The health effects caused by nickel exposure, described above, are mediated by active change in the expression of genes that control inflammation, the response to stress, cell proliferation, or cell death. Alteration in gene expression following nickel exposure is caused by the activation or suppression of a number of transcription factors, by changes in the levels of histone modifications and by alterations in DNA methylation. An important question is whether there is a specific pattern of gene expression in response to low, subtoxic doses of nickel. If such pattern does exist it could be utilized as a biomarker of nickel exposure. Nickel essentiality in mammalian cells is still debated. Therefore, no specific proteins involved in uptake, intracellular distribution or storage of nickel ions are known that might be coordinatively up- or downregulated by nickel exposure. Thus, a priori, it is difficult to predict that there are changes in gene expression that are characteristic to nickel exposure. Rather, one may assume that the expression of genes may be modified by the interaction of the nickel ions or reactive metabolites (e.g., reactive oxygen species) with elements of signal transduction cascades such as second messengers, protein kinases, phosphatases or transcription factors. Remarkably, nickel exposure produces a rather specific pattern of changes in gene expression, by inducing or suppressing genes coding for proteins involved in response to oxygen shortage (hypoxia) [23]. The mechanism involves nickel’s effect on cellular oxygen sensors. This type of response is better studied in an Met. Ions Life Sci. 2, 581–618 (2007)

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in vitro system, although some data indicate that it also exists in vivo. The activation of this set of genes has relevance to nickel carcinogenic activity. Another specific pattern of changes in gene expression is the activation of inflammatory response which is more easily noticed in vivo. The activation of genes coding for cytokines and chemokines correlates well with known nickel-induced allergic reactions and asthma. Undoubtedly, differences in cellular uptake and distribution of nickel ions in different organs may be manifested by variations in these patterns of gene expression. The activation of the hypoxic signaling pathway and switch of cellular metabolism to a state that mimics hypoxia may play a significant role in nickel-induced carcinogenesis [23,24]. Hypoxia has been shown to be common in tumors [25]. It may promote tumor progression via mechanisms enabling cells to overcome nutritive deprivation, to escape from the hostile metabolic microenvironment and to favor unrestricted growth [25]. The exposure of cells to nickel triggers cellular reactions typical of hypoxia, including the upregulation of expression of genes involved in glucose transport and glycolysis. Additionally, cellular responses to hypoxic stress include inhibition of cell proliferation and, when cell damage is irreversible, apoptosis. Therefore, nickel’s imitation of the state of hypoxia may provide the conditions for the selection of cells that have altered energy metabolism, changed growth control, and/or become resistant to apoptosis. Such selection requires time; however, since the switch to a ‘glycolytic phenotype’ in cells exposed to nickel lasts only as long as nickel is present, repetitive cycles of exposure are needed to facilitate the process. The selection theory seems to explain high transforming activity of nickel compounds, despite their low mutagenicity [26]. However, one may suggest that for successful cellular transformation an additional mutagenic (DNA damage) event is required. Indeed, nickel compounds are synergistic with many mutagenic carcinogens in enhancing cell transformation both in vitro and in vivo [27–29]. Additionally, the importance of epigenetic changes exerted by nickel compounds should be taken into consideration. The induction of cytosine methylation and histone methylation and deacetylation may lead to the inherited inactivation of the expression of senescence/tumor suppressor gene(s) and additionally contribute to the carcinogenic mechanism [30,31]. It is worthy emphasizing that the yield of nickel-induced tumors is known to be both tissue- and species-dependent (see also Kasprzak and Salnikow, Chapter 17). This suggests that genetic predispositions, including variations in the expression of genes involved in the metabolism of antioxidants (most likely glutathione and vitamin C) in different species and strains of animals, may also play an important role in nickel carcinogenesis [32]. It is conceivable that similar genetic predispositions take place in human populations. For years, changes in gene expression were analyzed by classic experimental approaches using Northern blotting or RT-PCR. More recently, a real-time PCR and DNA microarray techniques have emerged as assays that allow simultaneous Met. Ions Life Sci. 2, 581–618 (2007)


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screening of changes in a multitude of gene activities in one experiment. Such gene expression analyses have been done for nickel-exposed cells cultured in vitro and for exposed tissues of experimental animals and are discussed in Sections 3 and 7.

2. GENETIC AND EPIGENETIC CHANGES IN NICKEL-EXPOSED CELLS Nickel compounds are considered clastogenic, but only weakly mutagenic [33]. The clastogenic effect of nickel can be observed both in vivo and in vitro and in some cases it can be traced even after tumor formation. Thus, in one study, intramuscular injections of crystalline nickel sulfide resulted in the appearance of rhabdomyosarcomas in mice [34]. Three cell lines established from these tumors were characterized by a rearranged marker chromosome that was present in a majority of the cells of each line. The control tumors were obtained with the injection of the potent carcinogen, 3-methylcholanthrene. None of the 3-methylcholanthrene-induced cell lines contained rearranged marker chromosomes. The other study showed clastogenic activity of insoluble Ni3S2 by a clear increase in number of micronuclei in exposed human lymphocytes [35]. Both soluble and insoluble nickel compounds have been shown to generate specific chromosomal damage, particularly in the heterochromatic regions of the genome [36,37]. This effect has been especially noticeable in the heterochromatic long arm of the X chromosome, which suffers regional decondensation, frequent deletions and aberrations following exposure of cultured cells to insoluble NiS and soluble NiCl2 [36]. Similar chromosomal aberrations were also observed in nickel-transformed Chinese hamster embryo cells [38]. In cultured human lymphocytes, a maximal 1.9-fold induction of sister chromatid exchange (SCE) was detected following NiSO4 exposure [39]. In addition to chromosomal damage, DNA oxidative damage and the inhibition of nucleotide excision repair following nickel exposure were observed in mammalian cells in culture as well as in animal experiments [29,40–43]. In spite of these effects, mutagenesis assays revealed low mutagenic activity of nickel compounds in most of the mutational systems examined thus far, from Salmonella to mammalian cells in vitro [35,44–46]. In one study, a 2-h in vitro treatment of freshly isolated mouse nasal mucosa and lung cells with Ni3S2 clearly induced DNA fragmentation in a concentrationdependent manner. However, when a similar treatment was applied to lacZ and lacI Big Blue transgenic mice and rats, the mutation frequency of these target genes in the respiratory tissues was not increased [47]. Additionally, no increase of ouabain-resistant or 6-thioguanine–resistant colonies has been found in human diploid fibroblasts, even at concentrations of Ni3S2 which increased the frequency of anchorage-independence 200-fold [18]. No 6-thioguanine-resistant colonies Met. Ions Life Sci. 2, 581–618 (2007)

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were observed in hamster V79 cells exposed to two particulate nickel compounds, NiS or NiO black [46]. However, in the same study the G12 transgenic gpt cell line was very responsive to four insoluble nickel compounds, producing a 20- to 100-fold increase in the number of 6-thioguanine–resistant ‘mutants’. Christie et al. and others also have studied V79 cells transfected with pSV2gpt plasmid [48]. Insoluble nickel sulfide produced up to 80 times more ‘mutants’ over the spontaneous background level in one transgenic clone G12, but not in the parental cell line. It is noteworthy that the evidence of nickel mutagenesis in mammalian cells was mostly obtained using G12 and to a lesser extent G10 transgenic cell lines. Moreover, mutagenesis of the gpt target could not be demonstrated with other metals such as chromium, mercury and vanadium [49]. These ‘mutants’, as realized later, appeared to be silenced variants in which the transgenic gpt gene was inactivated by DNA methylation [30]. The mutagenic effect of NiCl2 was also studied using rat kidney cells infected with the murine sarcoma virus mutant ‘ts110’ [50]. The exposure of cells to NiCl2 produced a seven-fold increase in the reversion of the transformed phenotype in comparison with the spontaneous reversion frequency. As in G12, system changes in the expression mutations, leading to alternative RNA splicing were found in the transgene carrying DNA of the murine sarcoma virus mutant ts110. For example, one representative nickel-induced revertant clone had a mutation affecting MuSVts110 RNA splicing [51]. Other experiments with the SHE system have provided confirmatory evidence that cell immortalization can occur as an indirect consequence of carcinogen exposure following an induced high-frequency change in the treated population, rather than through direct targeted mutagenesis [26].

2.1. DNA Methylation and Transcription In DNA of higher eukaryotes the methylation of cytosine residues in CpG dinucleotides has been identified as an important modification that leads to the modulation of gene expression. In general, increased cytosine methylation represses transcription [52]. Exposure to nickel compounds increases the DNA methylation, leading to the inactivation of gpt gene expression [30]. The position of the gpt transgene on the chromosome was found to be an important factor, since the exposure of cells to nickel compounds resulted in hypermethylation of the transgene when it was located near the heterochromatin, but not when the transgene was located more distantly. Although the mechanisms by which nickel induces DNA hypermethylation are presently unknown, a possible model includes the ability of nickel to increase chromatin condensation and trigger de novo DNA methylation [30]. In vivo experiments demonstrated that the inactivation of a tumor suppressor gene can be associated with nickel-induced transformation. Thus, the injection of nickel sulfide into wild-type C57BL/6 mice as well as a mouse heterozygous Met. Ions Life Sci. 2, 581–618 (2007)


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for the tumor suppressor gene p53 produced malignant histiocytomas in all mice [53]. All tumors demonstrated hypermethylation of the promoter of the tumor suppressor gene p16. Identification of other tumor suppressor genes silenced by DNA methylation in Ni-exposed or transformed cells is in progress.

2.2. Histone Modification and Transcription In addition to gene silencing by DNA methylation the suppressive effect of nickel on histone H4 acetylation in vitro, in both yeast and mammalian cells, has been reported [31,54]. Acetylation of lysine 12 and 16 in histone H4 in yeast was more greatly affected than lysine 5 and 8, and it was proposed that nickel’s binding to histidine 18 in histone H4 may be responsible for this effect. Both the loss of histone acetylation and the increase in DNA methylation by nickel worked together in gpt gene silencing in the G12 transgenic cell line [55]. The acetylation of the core histone N-terminal ‘tail’ domains is now recognized as a highly conserved mechanism for regulating chromatin functional states. Biochemical data supports a correlation between histone acetylation, histone methylation, and gene expression, suggesting that histone modifications (histone code) act to enhance or decrease the access of transcription-associated proteins to DNA. Modulation of gene expression is important for nickel-induced transformation. Recent experiments showed that the exposure of nickel-transformed cells to the histone deacetylase inhibitor trichostatin A (TSA) resulted in the appearance of a significant number of revertants, measured by their inability to grow in soft agar [56]. Moreover, that study showed that treatment of cells with TSA inhibited the ability of nickel to transform mouse PW or human HOS cells to anchorageindependent growth. Taken together, these data point out that the epigenetic changes are likely to be more important for nickel-induced toxic and carcinogenic effects than mutational changes.

3.

ALTERATION OF GENE EXPRESSION FOLLOWING NICKEL-INDUCED LUNG INJURY

In recent years, a significant amount of work has been done to identify genes that control individual susceptibility to nickel-induced acute lung injury. The DNA microarray analysis of 8734 sequences allowed the examination of the responses of inbred mouse strains exposed to nickel sulfate. The changes in gene expression were related to increased oxidative stress, extracellular matrix repair, cell proliferation, and hypoxia, followed by a decrease in surfactantassociated proteins [57]. Certain expressed sequence tags (ESTs) were clustered with known genes, suggesting possible co-regulation and novel participants in pulmonary injury. Met. Ions Life Sci. 2, 581–618 (2007)

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Locus number estimation and a genome-wide analysis revealed that five genes could be responsible for the survival time and indicated significant linkage for a quantitative trait locus on chromosome 6, Aliq4 (acute lung injury QTL4). Haplotype analysis identified an allelic combination of five quantitative trait loci that could explain the difference in sensitivity to acute lung injury between parental strains. Positional candidate genes for Aliq4 include aquaporin-1 (Aqp1), surfactant protein B (SP-B), and transforming growth factor-α (TGF-α). These findings suggest that NiSO4-induced acute lung injury is a complex trait controlled by at least five genes (all possibly involved in cell proliferation, tissue regeneration, and surfactant function) [57].

3.1.

Genes Coding for Metal-Binding Proteins

Exposure to nickel sulfate (NiSO4) aerosols (110 µg/m3) for 3, 8, 24, 48, or 96 h causes acute lung injury in mice as evidenced by perivascular distension, epithelial damage, alveolar congestion, hemorrhage, neutrophilic infiltration, and pulmonary edema [17]. Differential analysis of changes in gene expression, which represents temporal and functional relationships of genes after pulmonary insult produced by NiSO4 exposure, has been done using microarray technique [58]. All analyzed genes were assigned to four groups: group I contained genes that tended to increase consistently throughout nickel-induced acute lung injury; group II contained genes that displayed a delayed increase; group III contained genes that tended to decrease throughout the response to injury; and group IV contained genes that displayed a delayed decrease. In group I the expression of two metal-binding proteins, metallothionein-1 (Mt-1) and lactotransferrin was consistently increased. The expression of Mt-1 was increased more than any other gene in that group (⬎ 13-fold at 96 h). Mt-1/2 are small-molecular-weight cysteine-rich proteins that preferentially bind zinc and to a lesser extent copper [59]. They have been widely studied for their role in the detoxification and homeostasis of essential and non-essential metals. Mt-1/2 may limit the toxicity of nickel by direct binding or through acting as an antioxidant with the capability to scavenging the reactive oxygen species. The role of Mt-1/2 in nickel-induced acute lung injury was evaluated using the Mt-transgenic as well as Mt1/2⫹/⫹ and Mt1/2⫺/⫺ mice [60]. Mt1/2⫺/⫺ mice were more susceptible than Mt1/2⫹/⫹ mice to nickel-induced inflammation, surfactantassociated protein B (SFTPB) transcript loss, and lethality. In contrast, Mttransgenic mice exhibited less lung injury and increased survival following nickel exposure. Thus, the expression of Mt-1/2 ultimately improves survival in the progression of nickel-induced acute lung injury in mice. The induction of Mt-1/2 also protected rat liver from the toxic effects of nickel following acute exposure [61]. However, Mt-1/2 overexpression did not prevent nickel-induced tumor formation in mice that received intramuscular injections of nickel [62,63]. Similar, to the Met. Ions Life Sci. 2, 581–618 (2007)


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in vivo response in cultured human lung epithelial cells, nickel exposure caused a dose-dependent increase of Mt-1 expression [64]. In our laboratory we also found an increase in Mt-1 expression in 1HAEo- and A549 human lung epithelial cells exposed to 0.25–0.5 mM NiSO4. Thus, the induction of Mt-1/2 expression seems to be a characteristic response to nickel exposure in lungs [65]. The induction of genes coding for two iron-binding proteins, ferritin and lactotransferrin, by nickel in human cultured lung cells, in mouse lung and in Chinese hamster ovary cells (CHO) cells [58,64,66] may reflect a disturbance in iron metabolism because of the ability of these proteins to sequester free iron [67–70]. However, this induction could also have a protective effect against oxidative stress since ferritin-L and -H genes have an antioxidant-response element, and therefore their expression is sensitive to the intracellular levels of antioxidants [71]. Another gene involved in iron metabolism, which was induced in cultured cells as well as in liver, lung, and brain of rats exposed to nickel, was heme oxygenase-1 (HO-1) [58,72]. HO-1 catalyzes the first and rate-limiting step in the degradation of heme to yield equimolar quantities of biliverdin IXa, carbon monoxide, and iron [73]. The ability of nickel to induce the enzyme in rat kidney was described previously [74,75]. In the microarray experiment, HO-1 displayed a delayed increase (group II) following nickel sulfate exposure in mice lungs [58]. Complexing nickel with sulfhydryl agents completely blocked the activation of HO-1 in vitro. Administrating cysteine orally prior to or shortly after administration of nickel had a similar blocking effect. As for Mt-1, the induction of HO-1 expression may be protective since elevated levels of HO-1 have been found to suppress inflammation and protect against hyperoxia- or hypoxia-induced acute lung injury [76,77]. Another gene, coding for iron-bound protein cytochrome Cyp2f2, displayed a delayed decline in expression (group IV) [58].

3.2. Surfactant Genes Expression As mentioned above, in experimental animals, exposure to NiSO4 aerosols results in acute injury to the lungs, which involves inflammation, extracellular matrix alterations, fibrinolysis, and disruption in surfactant homeostasis [57]. The microarray analysis showed that the expression of genes coding for surfactant proteins (SP) B and C was initially unchanged from high constitutive levels, but began to decline at 24 h and continued to diminish throughout the exposure (group IV) [58]. The expression of surfactants is extremely important for lung functions. They modulate alveolar surface tension, prevent atelectasis, inactivate reactive oxygen species, and augment host antimicrobial defenses [78,79]. Diminished expression of surfactants in lungs of nickel-exposed animals coincided with diminished expression of other genes involved in surfactant and phospholipid production, metabolism, and trafficking [80]. For instance, expression of phospholipids transfer protein (Pltp), which assists in the uptake of secreted Met. Ions Life Sci. 2, 581–618 (2007)

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surfactant lipids and plays a role in surfactant lipid trafficking and reutilization in alveolar type II epithelial cells, was suppressed by two- to three-fold. Expression of sodium phosphate transporter (Slc34a2), which recycles intracellular phosphate and is an essential component for surfactant phospholipid synthesis, was suppressed from six- to seven-fold. Expression of fatty acids synthase (Fasn) and stearoyl-coenzyme A desaturase 2 (Scd2), which produce important constituents of surfactant, was also significantly decreased by nickel exposure. In addition, expression of napsin A aspartic peptidase (Napsa), which is involved in the proper processing of surfactant-associated protein B and maturation of surfactants, was suppressed from two-and-a-half- to four-fold. Taken together, these data indicate that the expression levels of several essential genes in the synthesis and function of pulmonary surfactant are inhibited, and these expression changes may cumulatively lead to surfactant disruption in nickel-induced acute lung injury. Further work revealed that NiSO4 injury caused by diminished SP-B expression can be alleviated by increased expression of TGF-α [57,81]. Using the TGF-α transgenic mouse model, overexpression of TGF-α was shown to attenuate the inflammatory response, reduce pulmonary edema, and preserve levels of SP-B. TGF-α mRNA and protein are hypoxia-inducible and are overexpressed in VHL(–/–) renal clear carcinoma cells [82]. Nickel upregulates HIF-1 transcription factor and a significant number of hypoxia-inducible genes through a mechanism described in details in Section 6.1.1. It is clear that TGF-α production protects nickel-exposed lung cells similarly to that observed in VHL(–/–) renal clear carcinoma cells [82]. Activation of the HIF-dependent pathway by nickel induces also genes that may contribute to lung injury, nitric oxide synthase (iNOS) is one of them [83]. Indeed, the inhibition of nitric oxide synthesis during nickel exposure was shown to attenuate cytokine expression and inflammation, restore surfactant gene expression, and increase survival [84].

3.3. Genes Coding for Extracellular Matrix Proteins The extracellular matrix (ECM) is composed of a variety of molecules and includes numerous proteins of the collagen family, elastic fibers, glycosoaminoglycans and proteoglycans, and adhesive glycoproteins. The ECM plays an important role in maintaining structural integrity, and regulating cell polarity, migration, injury repair, and survival. Nickel exposure disrupts functioning of ECM by downregulating some collagens and vitronectin [85]. At the same time, the amount of TGF-β1 protein was increased in bronchoalveolar lavage (BAL) after exposure to nickel [80]. Since TGF-β1 is a multifunctional cytokine whose broad modulatory mechanisms affect numerous biological functions important to the synthesis and remodeling of the ECM, it was suggested that TGF-β1 plays an important role in the acute lung injury caused by nickel exposure [80]. Indeed, the activation of the TGF-β signaling pathway by nickel exposure results in the induction of expression of secreted Met. Ions Life Sci. 2, xx–xx (2007)


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phosphoprotein 1 (Spp1), a tissue inhibitor of metalloproteinase 1 (Timp1) and tenascin C (Tnc). Spp1 is a matricellular protein that possesses cytokine-like properties and can act as a chemoattractant for macrophages. Timp1 is a metalloproteinase inhibitor that can mediate inflammation and repair processes during acute lung injury through stabilization of matrix components. Tnc is an extracellular matrix glycoprotein that may play a role similar to Timp1. The expression of Spp1, Timp1, and Tnc was markedly increased after 72 h of nickel exposure, suggesting that the activation of the TGF-β pathway has an effect on the expression of extracellular matrix-altering genes during nickel-induced acute lung injury. In human keratinocytes (HaCat) nickel exposure induced the expression of matrix metalloproteinases 2 and 9 [86]. Matrix metalloproteinases are key enzymes involved in ECM degradation in physiological and pathological conditions. Additionally, genes coding for two enzymes involved in the hydroxylation of collagen, proline and lysine hydroxylase, are also significantly upregulated in nickel-exposed cells [24,87]. Hydroxyproline stabilizes the collagen triple helix under physiologic conditions [88]. Hydroxylysine participates in the formation of intermolecular cross-links, which impart mechanical stability to the collagen fibers.

3.4.

Genes Coding for Cytokines and Chemokines

The TGF-b1-dependent pathway is not the only one activated by nickel in lungs. The pathogenesis of lung injury may involve multiple signaling pathways: one that directs an insult on epithelial and alveolar cells (the consequences of this insult were described above) and others that involve the activation of alveolar macrophages in the lung, leukocyte recruitment and intercellular signaling between infiltrating leukocytes and the endothelial cells. Indeed, a number of chemokines and cytokines, such as IL-1b, Cxcl1 and CCl7, CCl9, CCl11, CCl17, and CCl22 have all been shown to increase after NiSO4 challenge in animal models [85]. One signaling pathway likely involves macrophage stimulating 1 receptor (Mst1r, a c-met-related tyrosine kinase, also known as the Ron receptor), which has been shown to play an important role in nickel-induced lung injury in mice [89]. Ron tk–/– mice succumb to nickel-induced lung injury earlier. They display larger and earlier increases in IL-6, CCL2, and macrophage inflammatory protein-2 expression. The serum of knockout Ron mice displays greater nitrite levels, indicating pulmonary pathology and augmented pulmonary tyrosine nitrosylation. Increases in cytokine expression and cellular nitration can lead to tissue damage and are consistent with the differences between genotypes in the early onset of pathology and mortality in Ron tk–/– mice. The upregulation of proinflammatory cytokines IL-6 and IL-8 induced by nickel was observed in numerous studies both in vitro and in vivo [67,86,90-93]. Met. Ions Life Sci. 2, 581–618 (2007)

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However, the underlying mechanism of upregulation of these cytokines by nickel still remains obscure. In one study, in BEAS-2B human airway epithelial cells, exposure to non-cytotoxic levels of Ni3S2 was shown to induce expression of IL-8, most likely via the induction of AP-1 and some other transcription factors [90]. A lung-infection-like mycoplasma could have a potentiating effect on IL-6 production following nickel exposure [94].

4.

NICKEL-INDUCED ALLERGY AND GENE EXPRESSION

Nickel-induced expression of cytokines and chemokines not only plays a central role in lung injury, but also is a key factor in nickel-caused allergic contact dermatitis [92,95]. Prostheses or other surgical implants made from nickel-containing alloys have been reported to cause nickel sensitization or to aggravate existing dermatitis. Although the elucidation of the molecular mechanisms of nickelinduced allergies has just begun, a rather detailed picture of cell interactions and activation of cytokine production has emerged [96]. According to these studies, nickel activates skin dendritic cells at the site of contact, promoting their migration to regional lymph nodes where T-cell priming occurs. Memory/effector T cells, due to the expression of skin-homing receptors, are rapidly recruited at the site of nickel challenge. Depending on the nickel doses, keratinocytes at the site of exposure may be targeted for apoptosis, mostly due to the intervention of nickel-specific CD8⫹ T cells. Type 1 chemokines (CCL1) released by activated CD4⫹ and CD8⫹ T cells activate keratinocytes and other resident cells, which in turn release IL-1α, IL-6, IL-8, TNF-α, GM-CSF, CCL2, CCL5, CCL20, CCL27, and some other cytokines and chemokines for the amplification of the inflammatory reaction [92,96]. CCL1 produced by keratinocytes, dendritic cells and activated T cells is critical for the recruitment of regulatory T-cell subsets (Tr1), which provide the feedback regulation through upregulation of expression and release of IL-10. IL-10 blocks the activation of T cells by impairing the antigenpresenting function of dendritic cells, thus limiting the extent of inflammatory response and tissue disruption. The suppressive function of Tr1 on dendritic cell function is strictly IL-10-dependent and can be reverted by anti-IL-10 antibodies [97]. In nickel-allergic individuals, Tr1 cells were found at a lower frequency [98]. Additionally, peripheral blood mononuclear cells (PBMC) from nickel-allergic individuals respond to Ni(II) with significantly greater production of interleukins IL-4, IL-5, IL-13, and INF-γ than PBMC from healthy individuals [99,100].

5.

NICKEL-INDUCED EXPRESSION OF ERYTHROPOIETIN

The unilateral intrarenal injection of nickel subsulfide (Ni3S2) produced pronounced erythrocytosis at one to four months after injection in rats or guinea pigs Met. Ions Life Sci. 2, 581–618 (2007)


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[101]. Erythrocytosis also occurred in rats after intrarenal implantation of Ni3S2 within semipermeable cellulose tubules, indicating that slow dissolution rather than phagocytosis of Ni3S2 particles is necessary for erythropoietic stimulation. The mechanism of erythropoietic stimulation include the induction of erythropoietin (Epo) expression which was upregulated five- to six-fold at two weeks after intrarenal injection of Ni3S2 [102,103]. Epo is the hormone that stimulates red blood cell proliferation and is synthesized in the kidney and liver normally in response to hypoxia. Similar to in vivo studies, exposure of human hepatoma cell line Hep3B to nickel chloride caused upregulation of Epo expression [104]. This increase in Epo expression in vitro and in exposed animals is clearly due to the activation of the HIF-1 transcription factor, since Epo expression is under the control of this transcription factor [105].

6. 6.1.

ALTERATION OF TRANSCRIPTION FACTORS AND SIGNALING PATHWAYS Protein Hydroxylation and Ascorbate

Protein hydroxylation plays an important role in protein–protein interactions. The best known example of these interactions is hydroxylation of proline and lysine residues in collagen molecules, which serves to form and stabilize the collagen triple helices. Collagen chains that do not contain 4-hydroxyproline cannot fold into triple-helical molecules that are stable at body temperature [88,106]. Another example of the role of hydroxylation in assisting protein– protein interactions is the hydroxylation of proline and asparagine residues in HIF-1α protein. The hydroxylation of proline residues 402 and 564 allows interaction of the HIF-1α protein with the von Hippel Lindau (VHL) protein. The hydroxylation of the asparagine 804 residue prevents interaction of the HIF-1α protein with acetyltransferase p300. The hydroxylation reaction is carried out by non-heme dioxygenases [107]. Among them are two long-known collagen prolyl-4-hydroxylases [108] and the more recently identified FIH-1, HPH1, HPH2, and HPH3 asparaginyl and prolyl hydroxylases, responsible for HIFα hydroxylation (Fig. 1) [109–112]. In the course of the hydroxylation reaction, the enzyme splits dioxygen into two atoms, one of which is converted into a hydroxyl group. Hydroxylases employ both iron and ascorbate as cofactors while utilizing 2oxoglutarate and O2 as cosubstrates, each of which could be a potential target for the regulation of activity of these enzymes. The role of ascorbate is to maintain iron as Fe(II), which makes the enzyme functional. The exposure of cells to nickel compounds depletes intracellular ascorbate both by blocking ascorbate entry into cells, via the inhibition of expression of sodium-dependent vitamin C transporter (SVCT2), and by catalizing ascorbate oxidation inside and outside the Met. Ions Life Sci. 2, 581–618 (2007)

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Figure 1. Effect of nickel exposure on protein hydroxylation. The hydroxylation of proline, lysine, and asparagine molecules is carried out by non-heme dioxygenases. Among them are two collagen prolyl-4-hydroxylases (C-P4H 1 and 2) and the FIH-1, HPH1, HPH2, and HPH3, asparaginyl and prolyl hydroxylases, respectively responsible for HIF-1α and HIF-2α hydroxylation. Hydroxylases utilize 2-oxoglutarate and O2 as co-substrates and ascorbate as a cofactor, each of which could be a potential target for regulation activity of these enzymes. The role of ascorbate is to maintain iron as Fe(II), which makes the enzyme active. The asparagine hydroxylation, which is mediated by FIH protein, prevents complex formation between HIF-1α and the CBP and p300 transcriptional co-activators. Hydroxylation of proline residues 402 and 564 in the oxygen-dependent degradation domain (ODD) of HIF-1α leads to its interaction with the VHL tumor suppressor protein, a part of the ubiquitin– ligase complex. This results in ubiquitylation and rapid proteasomal degradation of HIF-α proteins. The exposure of cells to nickel compounds depletes intracellular ascorbate. This results in the inactivation of the hydroxylases, stabilization of HIF-α proteins, and formation of the HIF-1 transcription complex. Thus, exposure to nickel produces a phenotype observed in hypoxic cells or in cells with mutated VHL. Loss of C-P4H1 and C-P4H2 activities prevents formation of collagen triple helical fibers and leads to disorganization of ECM.

cell [32,113]. This results in inactivation of the hydroxylases and produces a phenotype observed in hypoxic cells or in cells with mutated VHL [32,113]. The addition of 100 µM ascorbate to the culture medium restored intracellular ascorbate levels in nickel-exposed cells [113]. The elevation of intracellular ascorbate level Met. Ions Life Sci. 2, 581–618 (2007)


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correlated well with activation of HIF-1α hydroxylation and decrease of HIF-1dependent transcription. On the other hand, exposure to nickel can disturb iron metabolism since nickel may compete with iron for the same divalent metal transporter (DMT-1) [114]. The chelation of hydroxylase-bound iron by chelators is known to inhibit the enzymes and produces hypoxic response [115]. Thus, the likelihood exists that any alteration of iron metabolism could also affect the PHD or FIH-1 enzymes. Since Ni(II) is similar to Fe(II), it was suggested that the induction of the hypoxic response may occur by direct substitution of Ni(II) for Fe(II) at the active site of the hydroxylases. While the possibility of metal substitution for these enzymes in test-tube experiments has been suggested [115], the evidence is still lacking for such substitution in cells. Furthermore, metal ion movement within the cell is highly regulated, with most ions bound to either small carrier molecules or protein chaperones. Thus, the data that have been presented for a model in which the addition of nickel inhibits HIF hydroxylases by depleting the cellular stores of ascorbate [32,113] more likely represent the mechanism of HIF induction by nickel. While other data are in agreement with a model of ascorbate depletion, finer details of the precise amount and fluctuation of ascorbate in vivo could lend additional support [116]. Thus, similar to hypoxia, the exposure of cells to nickel, results in the induction of the HIF-1 transcription factor and activates the expression of hypoxia-inducible genes [23].

6.1.1.

HIFa Hydroxylation and Transcriptional Response

The HIF-1 transcription factor is a HIF-1α /HIF-1β (ARNT) heterodimer, which is formed in response to low oxygen tension in the cells. Under hypoxic conditions, it activates transcription of a number of target genes whose promoters contain the binding motif termed the ‘hypoxia-response element’ (HRE) [23]. The α subunit is the regulatory component of the HIF-1 complex and is unique to the hypoxic response (Figure 1). Under normoxic (normal oxygen level) conditions, this protein is virtually undetectable in most cells due to hydroxylation and rapid proteasomal destruction, but can accumulate following exposure to proteasomal inhibitors such as lactacystin or MG-132 (Figure 1) [117]. Accumulation of the HIFα subunit in the presence of hypoxia or nickel implies that proteasomal degradation of the protein is impaired by these exposures. A molecular basis for the HIF-1α degradation under normoxic conditions and stabilization under hypoxic conditions is now known in detail. Hydroxylation of proline residues 402 and 564 in the oxygen-dependent degradation domain of HIF-1α leads to its interaction with the VHL tumor suppressor protein, a part of the ubiquitin–ligase complex (Figure 1) [118–120], followed by ubiquitylation and rapid proteasomal degradation of HIF-1α. The foundation for specific protein–protein interaction is provided by the introduction of the hydroxyl group Met. Ions Life Sci. 2, 581–618 (2007)

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at the proline 4 position, where the oxygen of this hydroxyl group facilitates hydrogen bonding with VHL’s Ser-111 and His-115 residues. This is sufficient to enable discrimination between non-hydroxylated and hydroxylated HIF-1α protein [119–123]. Besides proline, hydroxylation of asparagine residues was found to be important for HIF-α protein interactions. Thus, asparagine hydroxylation, which is mediated by FIH protein, prevents a complex formation between HIF-1α and the CBP and p300 transcriptional co-activators which by this way also become involved in the control of HIF-dependent responses [112,124]. The upregulation of HIF-1α protein following nickel exposure was first described by Salnikow et al. in human cell lines including A549, MCF-7 and HOS cells [125]. Significant amounts of HIF-1α protein were observed in A549 cells after 6 h of nickel exposure, and the HIF-1α protein level remained high after 24 h. In agreement with the presented mechanism of HIF-1α protein upregulation by hypoxia, nickel also upregulated the HIF-1α protein via the loss of proline hydroxylation [113]. The induced HIF-1 transcription factor upregulated transcription of reporters with HRE originated from Epo and iNOS [125]. Further work revealed that the induction of the previously cloned nickel-inducible gene NDRG1/Cap43 [126] by nickel is mediated by activation of the HIF-1 transcription factor [127]. In the next study, mouse fibroblasts originated from wild-type or HIF-1α knockout (HIF-1α⫺/⫺) mice, were used to identify genes induced or suppressed by nickel in a HIF-1-dependent manner [24]. Using the GeneChip technique combined with Northern blot and RT-PCR it was found that after NiCl2 treatment, 114 genes were upregulated and 66 genes downregulated over fourfold in a HIF-1-dependent manner, while 29 genes were upregulated and 31 genes downregulated over four-fold in a HIF-1-independent manner [24]. Thus, nearly 75% of gene expression induced by nickel and more than 50% of gene expression suppressed by nickel were HIF-1-dependent. The number of genes that were upregulated and downregulated by nickel treatment in HIF-1α-deficient cells was four times more than that in HIF-1α-proficient cells. Many genes, including NGFβ, SGK, IP10, CD44, heparin-binding EGF-like, melanocortin 1 receptor, Grg1, BCL-2-like and tubulin-binding protein E-protein EMap-115, were induced by nickel only in HIF-1α-deficient cells. It was suggested that the larger number of genes induced by nickel in HIF-1α-deficient cells may compensate for the loss of the HIF-1 and the transcription of downstream genes. This indicates that the loss of the normal response to hypoxia-like stress may lead to the activation of other mechanisms facilitating cell survival. The data emphasize the important role of HIF-1 in the cellular response to hypoxia, nickel or other hypoxia-mimicking agents. The comparison of HIF-1-dependent genes induced/suppressed by soluble and insoluble nickel compounds using GeneChip analysis revealed similar patterns, suggesting that the same mechanism is activated by different nickel compounds [128]. The important finding in GeneChip analyses was HIF-1-dependent induction by nickel genes coding for glycolytic Met. Ions Life Sci. 2, 581–618 (2007)


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enzymes and genes involved in glucose transport [24,129]. Of 12 candidate genes involved in glucose transport and glycolysis, 10 were induced by nickel exposure in HIF-1α-proficient cells, but not in HIF-1α-deficient cells. Glucose-6-phosphate dehydrogenase and hexokinase I, whose expression was reported not to be significantly affected by hypoxia, were also not significantly changed by nickel. In contrast, hexokinase II, that is known to be strongly stimulated by hypoxia, was induced more than 10-fold by nickel exposure in a HIF-1-dependent manner [24,129]. Moreover, glyceraldehyde-3-phosphate dehydrogenase (GAPDH) that can be induced by hypoxia or nickel was also found to be induced approximately 4.5-fold in HIF-1α-proficient cells, but not in HIF-1α-deficient cells. Besides glycolytic enzymes and glucose transporters GeneChip microarrays identified a significant number of other genes upregulated by nickel via activation of the HIF-1 transcription factor. These include genes encoding adrenomedullin, BNIP-3, carbonic anhydrase IX, ceruloplasmin, Dec1/STRA13, Epo, inducible nitric-oxide synthase, IGF-II, heme oxygenase-1, NDRG-1/Cap43, plasminogen activator inhibitor I, prolyl-4 hydroxylase, tyrosine hydroxylase, vascular endothelial growth factor (VEGF), and many others [24,64,87,104,127–133]. As mentioned earlier, not all genes were upregulated by nickel. Many genes were found to be significantly suppressed by nickel in a HIF-1-dependent manner. Monocyte chemoattractant protein 1 (MCP-1), which contributes to macrophage infiltration in human ovarian carcinomas, was found to be suppressed by nickel over 20-fold [24]. Among other down-regulated genes that may play an important role in mediating carcinogenic effects of nickel is Zac-1, a tumor suppressor gene that was found suppressed by nickel exposure only in HIF1α-proficient cells. Zac-1 is a zinc-finger nuclear transcription factor, which possesses antiproliferative effects and is frequently silenced in ovarian and breast cancer cells [134]. Other downregulated genes include VEGF-related transmembrane receptor neuropilin-1 (Npn-1), basement membrane protein laminin-5γ 2, and the growth-arrest-specific gene Gas-1, which may be a tumor suppressor. Gas-1 is distinct among them, since it could be downregulated in both HIF-1αproficient and HIF-1α-deficient mouse fibroblasts [24]. The nickel-induced suppression of tumor suppressors Zac-1 or Gas 1 may be an important part of the cell transformation process initiated by nickel exposure. It was suggested that HIF-1-dependent suppression of gene expression may be considered as a kind of metabolic adaptation, which facilitates a rescue process for the exposed cells [24]. This adaptation causes a persistent downregulation of cellular metabolism, in order to keep up with energy demand and supply throughout the hypoxic period. Similar conditions can be mimicked by nickel exposure; thus, nickel-induced suppression of gene expression is probably a part of a survival process. A recent study indicated that another gene, serpin, is subjected to downregulation by nickel [135]. Serpina3g, a member of serpin family, was the most downregulated gene in response to nickel exposure of mouse cells. A HIF-dependent Met. Ions Life Sci. 2, 581–618 (2007)

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mechanism is responsible for the downregulation of Serpina3g. A similar downregulation of Serpina3g transcription was found to be induced by hypoxia and other hypoxia mimetics, including cobalt, deferoxamine, and the proline hydroxylase inhibitor dimethyloxalylglycine (DMOG) [135]. The silencing was found to be a dynamic and reversible process because of the recovery of Serpina3g gene expression after removal of nickel. Moreover, the expression of nickel-silenced Serpina3g can be reactivated by TSA and 5-azacytidine, indicating that silencing by nickel resulted from either a direct or an indirect epigenetic mechanism [135]. The β subunit (ARNT) of HIF-1 is constitutively expressed and also is involved in xenobiotic responses where HIF-1β forms a dimer with the aryl hydrocarbon receptor (AhR) [105]. It is noteworthy that while ARNT expression itself is not directly affected by nickel, the expression of AhR-dependent genes can be significantly suppressed by nickel exposure [136]. The downregulation of AhR-dependent genes has an important toxicological implication. It may lead to a decrease in toxicant removal. Since cross-talk between the AhR-dependent pathway and HIF-dependent pathway has been described, it is conceivable that nickel affects a major regulatory factor upstream of both pathways. Indeed, the similarity in effects of nickel, hypoxia, and the inhibitor of 2OG-dependent dioxygenases, DMOG, suggests that iron and 2-oxoglutarate-dependent dioxygenases are involved in regulation of both pathways.

6.1.2.

Extracellular Matrix and Transcription

Focal adhesions are highly ordered assemblies of transmembrane receptors, extracellular matrix proteins, and a large number of cytoplasmic proteins, including structural proteins, as well as tyrosine kinases, phosphatases, and their substrates. They give rise to signaling platforms at the adhesive sites. The exposure of cells to toxic metals typically results in retraction and detachment from the surface. Nevertheless, HUVEC cells exposed to nickel remained on the surface without signs of retraction [137], but were unable to reach confluency, which was determined by the decrease in cadherin-5 expression. Despite this apparent cytotoxicity produced by nickel, focal adhesion distribution as visualized by immunofluorescence staining of vinculin was not affected [137]. Integrin-mediated interactions of cells with components of the extracellular matrix regulate cell survival, cell proliferation, cell differentiation, and cell migration. Some of these physiological responses are regulated via activation of transcription factors such as AP-1, EGR-1, NF-κ B. Exposure to 1 mM NiCl2 induces mRNA expression of E-selectin, intercellular adhesion molecule-1, (ICAM-1), vascular cell adhesion molecule-1 (VCAM-1), IL-6 and IL-8 [138– 141], whereas exposure to other metals like ZnCl2 and CrCl3 had no effect [138]. The induction of expression of surface molecules by nickel was through direct activation of transcription factors and not through an autocrine IL-1-dependent Met. Ions Life Sci. 2, 581–618 (2007)


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mechanism, which normally leads to the activation of expression of ICAM-1, VCAM-1 and E-selectin [141]. Since the transcription factor NF-κ B is known to be involved in the regulation of ICAM expression in endothelial cells, its response to nickel exposure has been investigated [138,142]. Treatment of HUVEC with nickel induced the translocation of NF-κ B p65 and also p50 subunit into the nucleus. NF-κ B binding activity was enhanced following nickel exposure, as determined by the DNA mobility shift analysis [138]. The activation of NF-κ B p65 at the protein level was accompanied by induction of mRNA expression for this gene. In addition to the enhanced DNA-binding activity of NF-κ B, another transcription factor, AP-1, was also augmented in HUVEC stimulated by nickel or by proinflammatory mediators and the phorbol ester (PMA) [138].

6.2. ATF-1 Exposure of cells to nickel induces a proangiogenic VEGF through activation of both the HIF-1 transcription factor and the AP-1 transcription factor [131]. At the same, time nickel is very efficient at turning off the expression of thrombospondin I (TSP I), an extracellular matrix protein [143,144]. The TSP I protein is a known regulator of angiogenesis in vivo [145]. High levels of this protein suppress growth of blood vessels into the tumor body. Thus, this is an excellent example of dual efficiency of carcinogenic nickel. The induction of VEGF and the loss of TSP I expression in nickel-induced tumors promote angiogenesis, thereby facilitating tumor growth. In vitro in nickel-transformed rodent cells the TSP I gene was found to be transcriptionally downregulated [143]. The TSP I gene appeared to be intact, based upon restriction enzyme analysis, and no DNA methylation changes were observed in the promoter of the gene [144]. It was found, however, that the ATF-1 transcription factor was hyperactivated in nickel-transformed cells and played the role of a negative regulator of TSP I [144]. The ATF-1 transcription factor belongs to an ATF/CREB family that was originally identified as a target of the cAMP signaling pathway [146]. Elevation of intracellular calcium by nickel (described in Section 6.4) also may activate a protein kinase cascade that mediated ATF/CREB phosphorylation. Thus, these data indicate that one or both of these pathways may be modulated in nickel-exposed and transformed cells.

6.3. Nickel-Induced Oxidative Stress Nickel complexes with certain natural ligands may be redox active at physiological pH although to a lesser extent than iron or copper complexes [147]. Nevertheless, they are capable of depleting important intracellular antioxidants such as Met. Ions Life Sci. 2, 581–618 (2007)

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glutathione and ascorbate [93,113,148]. This notion is supported by the finding that nickel resistance can be conferred by elevating intracellular levels of glutathione [148]. The exhaustion of reducing potential of the cell can be measured by enhanced DCFH fluorescence [149,150]. This oxidative stress may contribute to oxidative damage of proteins and DNA and activation of AP-1, NF-κ B, and other oxidatively sensitive transcription factors [148]. Nickel sulfides display higher carcinogenic activity than other derivatives of nickel, possibly because of an elevated production of reactive oxygen species (ROS) by these compounds [151]. More details on this subject can be found in Chapter 17. Generally, two separate mechanisms may be proposed for Ni(II) toxicity. One is direct, resulting from intracellular Ni(II) effects on reducing agents or reactions with endogenous oxidants, such as oxygen or H2O2, which increase the yield of the damaging ROS. The other mechanism, possible only in animals, is indirect and involves nickel-induced inflammation at the site of exposure and generation of ROS by inflammatory cells [152]. Nickel-resistant cells represent a good model for investigation of gene expression in cells chronically exposed to nickel. When comparative analysis of gene expression in parental and nickel-resistant cells has been done, the upregulation of several genes related to oxidative stress has been noticed [153]. Among these genes were COX-1 and COX-2. ROS-mediated upregulation of COX-2 was reported previously in cadmium [154] and arsenic-treated cells [155]. Another group of genes overexpressed in nickel-resistant cells was associated with glutathione metabolism. Thus, glutathione-S-transferase (GST) alpha was induced 2.2-fold, and theta 5.4-fold, while glutathione synthetase was induced 2.9-fold. These data provide an explanation for the high level of glutathione metabolism reported earlier for these cells [148]. Changes in GST activity were observed previously in mice treated with nickel [156,157]. Nickel treatment also resulted in a significant increase in lipid peroxidation and enzymatic activities of catalase and glutathione-S-transferase in vivo [61,157]. High levels of alanine aminotranferase and aspartate aminotransferase were found in serum of exposed animals. In contrast, the levels of reduced glutathione and superoxide dismutase were decreased. Interestingly, Zn(II) supplementation protected the activities of catalase, glutathione-S-transferase, the levels of GSH and decreased lipid peroxidation in nickel-treated rats.

6.3.1.

NF-lB

The heterodimeric protein NF-κ B (the predominant NF-κ B dimers are p65:p50) is a ubiquitous redox-regulated transcription factor that remains sequestered in the cytoplasm as an inactive complex with its inhibitory counterpart Iκ B [158]. Exposure to oxidative and inflammatory stimuli, such as TNF-α, IL-1, phorbol ester, ultraviolet radiation or microbial infection, leads to phosphorylation and Met. Ions Life Sci. 2, 581–618 (2007)


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subsequent proteasomal degradation of Iκ B, thereby releasing free NF-κ B dimers for translocation to the nucleus where NF-κ B binds to the promoter region of its specific target genes. Using electrophoretic mobility shift assays, a strong increase of NF-κ B DNA binding was detected upon stimulation of HUVEC cells with NiCl2 or CoCl2 [159]. These data were confirmed in human skin dendritic cells [160]. Among NF-κ B downstream genes are ICAM-1, VCAM-1, and endothelial leukocyte adhesion molecule-1 (ELAM-1, E-selectin). They play an important role in leukocyte recruitment to sites of inflammation during contact hypersensitivity. ICAM-1, VCAM-1, and ELAM-1 were found to be upregulated by NiCl2 exposure in cultured HUVEC cells [139]. This induction by NiCl2 required de novo mRNA and protein synthesis. Upregulation could be blocked by kinase inhibitor H-7, but not by staurosporine, suggesting involvement of phosphorylation events independent of protein kinase C activation. In normal human keratinocytes watersoluble nickel salts, nickel gluconate, nickel sulfate, and nickel chloride, upregulated production of ICAM-1, TNF-α and very late antigen-3 (VLA-3) [161]. Moreover, pretreatment for 24 h with NiCl2 produced hyporesponsiveness to IL-1 and TNF-α upon re-stimulation, suggesting that NiCl2 and these cytokines may partially share a common pathway of activation [139]. Another example of nickel-induced activation of a common pathway with TNF-α is the induction of MCP-1 messenger RNA and protein expression in endothelial cells [162]. Furthermore, NiCl2 was found to induce in a dosedependent manner mRNA production and protein secretion of proinflammatory cytokine IL-6. When the transcriptional mechanisms underlying gene-inductive effects of nickel were studied, the NF-κ B transcription factor was found to be involved in the inducible expression of adhesion molecules, IL-6 and MCP-1 [159,162,163]. NF-κ B is an important transcription factor in apoptosis and inflammatory response. It is clear that activation of NF-κ B by nickel causes significant modulation of cellular and tissue responses. Since the adhesion molecules, and numerous cytokines, and chemokines are under transcriptional control of NF-κ B, the activation of NF-κ B may explain nickel-induced allergic effects and contact skin hypersensitivity described in industrialized countries.

6.3.2. AP-1 AP-1 is a dimeric transcription factor composed of proteins that belong to the Jun (c-Jun, JunB, JunD), Fos (FosB, Fra-1, Fra-2), Maf, and ATF subfamilies [164]. The activation of AP-1 results from heterodimerization of proteins of Fos and Jun subfamilies. Formation of the proper transcriptional complex is important for the induction of genes which will respond appropriately to environmental stimuli including, but not limited to, oxidative stress. AP-1 activity is regulated by the redox state of a specific cysteine located at the interface between the two Met. Ions Life Sci. 2, 581–618 (2007)

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subunits, highlighting the importance of redox status on regulation of gene transcription and carcinogenesis [165]. AP-1 can be induced in response to certain metals, including nickel, H2O2, as well as several cytokines and other physical and chemical insults [166]. Expression of JunB, a subunit of AP-1, was found significantly up-regulated following nickel exposure [24,128]. In human airway epithelial cells BEAS-2B the addition of nickel induced both c-Jun and c-Fos mRNA levels [90,167]. The changes in gene expression were concomitant with the increase phospho- and total c-Jun protein levels and resulted in activation of AP-1 and downstream fibrotic genes, such as PAI-1 or IL-8. However, in these cells AP-1 induction seems to be oxidative stress-independent. Other studies reported nickel-stimulated activation of protein kinases that may be involved in the activation of AP-1, including p44/42 extracellular signalregulated kinases, p38, and stress-activated protein kinase/c-jun N-terminal kinases [168]. Thus, both ROS-dependent and ROS-independent pathways may play a role in activation of AP-1 in nickel-exposed cells. Following nickel exposure AP-1 could cooperate with other transcription factors, especially HIF-1 to maximize the induction of gene expression [90,131,132,167].

6.4. Nickel-Induced Changes in Calcium Homeostasis The role of changes in calcium homeostasis in cell transformation is not well understood. Ca2⫹ is recognized as one of the most important intracellular second messengers and is maintained at a very steep gradient between the outside and the inside of all mammalian cells [169]. Intracellular cytoplasmic levels of Ca2⫹ have been shown to signal gene expression changes associated with cell growth, differentiation, and apoptosis of many different types of cells in the body [169, 170]. Few studies have related carcinogenesis with the disturbances in calcium metabolism [171]. It can not be excluded that carcinogenic and/or toxic effects of nickel are also mediated by changes in calcium homeostasis. Thus, elevation in calcium concentrations in mice pancreas after nickel administration has been observed [172]. This elevation of calcium concentrations caused the activation of trypsinogen which resulted in the increase in trypsin activity in the tissue. One of the earliest observations that nickel-transformed cells displayed an unusual ability to rapidly proliferate in vitro in a low-calcium media suggested an alteration of intracellular calcium metabolism in cells originated from malignant rhabdomyosarcomas induced by Ni3S2 [173,174]. In vivo the addition of exogenous Ca2⫹ prevented the formation of lung adenomas caused by nickel or lead [175]. However, when added alone Ca2⫹ elevated the incidence of lung adenomas. Using NDRG1/Cap43 gene expression as a marker, we found that NDRG1/Cap43 was induced to the same extent by nickel or by the Ca2⫹ ionophore A23187, and that induction of the gene by nickel or the Ca2⫹ ionophore was abolished when Met. Ions Life Sci. 2, 581–618 (2007)


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free intracellular Ca2⫹ was chelated with 1,2-bis-(o-aminophenoxy)-ethaneN,N,N′,N′-tetra-acetic acid, tetra-acetoxymethyl ester (BAPTA-AM). The direct measurements confirmed the notion that free intracellular Ca2⫹ levels were elevated in nickel-treated cells [176]. The mechanisms leading to this elevation in intracellular Ca2⫹ in nickel-treated cells are currently not well understood. Furthermore, it is also not clear why NDRG1/Cap43 gene/protein expression is increased in tandem with these elevations of intracellular Ca2⫹. Soluble nickel enters the cell via DMT-1 transporter [114], but may use calcium channels since the calcium ionophore ionomycin (3 µM) increases nickel uptake four- to five-fold [177,178]. Additionally, nickel uptake into IHKE cells was inhibited by calcium. Ni2⫹ is known to be a T-type Ca2⫹ channel blocker [179,180], and it is conceivable that an initial decrease in intracellular Ca2⫹ level in response to the Ni2⫹ treatment was followed by a compensatory increase in free Ca2⫹ that resulted from its release from intracellular stores. In fact, nickel was found to evoke the release of stored intracellular calcium via a mechanism involving a cell surface receptor [181]. Another possibility is that Ni2⫹ ions interact with a Ca2⫹-sensor or a Ca2⫹-receptor on the plasma membrane to activate intracellular Ca2⫹ release. However, modulation of extracellular Ca2⫹ levels from zero up to 7 mM affected neither the expression of the calcium- and nickel-inducible gene NDRG1/Cap43, nor NDRG1/Cap43 induction by 1 mM NiCl2 [176]. The above results indicated that if Ni2⫹ ions were interacting with a surface Ca2⫹ receptor, it probably was not a Ca2⫹-binding domain, since it is unlikely that 1 mM NiCl2 would compete with 7 mM CaCl2 for binding at a specific protein site. Taken together, these data indicate that changes in calcium homeostasis invoked by nickel exposure may help in activation of gene expression induced via other signaling pathways. A good example is the evidence that activation of NDRG1/Cap43 and VEGF gene expression is mediated by combined interactions of HIF-1 transcription factor and calcium-induced protein kinases [131].

6.4.1.

NF-AT

NF-AT was originally described as a transcription factor expressed in activated T cells [182]. The activation of NF-AT proteins follows precisely the activation of calcineurin, the calcium/calmodulin-dependent phosphatase. In T cells, if intracellular free calcium levels remain elevated, calcineurin activity remains high, and NF-AT remains activated and nuclear-bound for many hours. NF-AT proteins are capable of binding cooperatively with transcription factors of the AP-1 (Fos/Jun) family to composite NF-AT:AP-1 sites, found in the regulatory regions of many genes that are inducibly transcribed by the immune system cells [182]. Numerous cytokines are under transcriptional control of NF-AT, including IL-1, IL-2, IL-3, IL-4, IL-5, IL-6, IL-8, IL-10, IL-13, IL-18, TNF- a, IFN-c, and GM-CSF [182]. Soluble NiCl2 or particulate Ni3S2, both markedly activated NF-AT in PW mouse fibroblast cells [183]. Met. Ions Life Sci. 2, 581–618 (2007)

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The elevation of intracellular calcium is likely to initiate the process of NF-AT activation, since pretreatment of cells with either nifedipine or BAPTA-AM resulted in an inhibition of NF-AT activation induced by Ni3S2 or NiCl2. Moreover, co-treatment of cells with thapsigargin, a cell-permeating chemical that induces the release of intracellularly stored Ca2⫹, and nickel had significant synergistic effects on NF-AT activation. Authors had speculated that Ca2⫹ release from stores is mediated by H2O2 produced following exposure to nickel compounds [183]. The induction of NF-AT could be responsible for activation of cytokines during nickel-induced inflammatory response.

6.5. PI3 Kinase and Downstream Transcription Factors The phosphatidylinositol-3-kinase (PI3K) signaling pathway is crucial to many aspects of cell growth and survival. The PI3K family constitutes a large family of lipid and serine/threonine kinases, which includes a number of phosphatidylinositol kinases, as well as the related DNA-dependent protein kinases. The induction of PI3K by nickel compounds is controversial. The induction has been observed by Li et al. in mouse epidermal Cl41 cells [184]. It was suggested that the activation of PI-3K/Akt-dependent pathway results in HIF-1 transactivation and NDRG1/Cap43 gene expression [184]. However, in other studies using A549 or BEAS-2B human lung epithelial cell the activation of HIF-1 and downstream genes NDRG1/Cap43 and PAI-1 expression by nickel was found to be not sensitive to wortmannin, a specific inhibitor of PI3 kinase [132, 176]. Moreover, in mouse epidermal Cl41 cells, the activation of PI3 kinase was shown to result in AP-1 induction [185]; yet no AP-1 induction following nickel exposure was observed in these cells [163].

6.6. DNA-Damage Response and Apoptosis Nickel-induced apoptosis was first reported in CHO cells [186]. However, in peripheral blood mononuclear cells, it was shown that high concentrations of nickel produced cell necrosis, whereas lower concentration caused apoptotic phenomena [187]. In primary cultures of rat hepatocytes nickel almost exclusively induced necrosis at 500 µM with very few cells undergoing apoptosis [188]. Therefore, the exposure to toxic concentrations of nickel results in cellular death through both necrosis and apoptosis. If death is caused by necrosis, then a strong inflammatory tissue reaction is likely to occur with the induction of cyto- and chemokines described above. The lower concentrations of nickel induce toxicity characterized by apoptotic phenomena, which does not produce inflammatory reactions, but induces apoptosis-specific genes. Indeed, low concentrations of nickel(II) acetate concentrations rapidly induce apoptosis in T-cell hybridoma cells [189]. Met. Ions Life Sci. 2, 581–618 (2007)


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In these cells, there was an increase in FasL protein levels and transient activation of caspase-3, a downstream effector of Fas/FasL-induced apoptosis. In another study, the treatment of human hepatoma Hep3B cells with nickel resulted in cell proliferation arrest, the appearance of detached cells, condensed chromatin, apoptotic bodies, and specific DNA fragmentation, indicating the occurrence of cell apoptosis [190]. This treatment inhibited expression of antiapoptotic Bcl-2. In this case the decrease of histone H4 acetylation in nucleosomes associated with the Bcl-2 promoter region was shown to be the cause of Bcl-2 downregulation. Further studies showed that increasing histone acetylation by either TSA or overexpressing histone acetyltranferase p300 in Hep3B cells attenuated the Bcl-2 downregulation and cell apoptosis caused by nickel [190]. Considering the importance of Bcl-2 in determining cell survival and apoptosis, the data presented here suggest that histone hypoacetylation may represent an unrevealed pathway in Ni(II)-induced cell apoptosis, where Bcl-2 is one of its targets. Additionally, strong induction of proapoptotic Bcl-2-binding protein BNIP3 by nickel has been reported [129]. Since BNIP3 is a HIF-1-dependent gene its induction by nickel is clearly the result of HIF-1 activation.

6.6.1.

P53

P53 can be described as a stress response gene [191]. Its product, the p53 protein is stabilized in response to DNA damage and acts to induce apoptosis or cell cycle arrest, thereby maintaining genetic stability of the cell. These functions are realized by a series of steps known as the ‘p53 pathway’, involving induction of the expression of a number of other genes [191]. Exposure to nickel may cause DNA damage possibly via the induction of oxidative stress. Indeed, accumulation of p53 protein following nickel exposure was reported [125]. Microarray analysis revealed changes in expression of genes activated by DNA damage caused by Ni(II) in cells, including Rad23 UV excision repair protein homolog and Gadd45 (growth arrest and DNA damage) [24,153]. Both genes take part in DNA repair. Cyclin kinase inhibitor p21 was also induced by nickel exposure in a number of cell lines [128,129]. Both Gadd45 and p21 genes are known to be transcriptionally activated by wild-type p53 protein following DNA damage, which causes cell cycle arrest at the G2/M phase. P53 is the most commonly mutated gene in human cancer. The p53 gene was reported to be mutated in human kidney epithelial cells chronically exposed to and eventually transformed by nickel [192]. However, no mutations in the p53 gene were found in analysis of ten nickel-induced rat renal tumors [193]. These results disputed the possible involvement of p53 mutations in nickel-induced transformation. Additionally, human HOS cells have mutant p53 [194], but nickel treatment results in their further transformation [21,95,196]. The acute Met. Ions Life Sci. 2, 581–618 (2007)

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treatment of human cells with NiCl2 does not change the expression for the p53 gene; however, it induces a wild-type p53 protein, but not mutant p53 [125]. Another report suggested the induction of p53 protein by nickel acetate in CHO cells [186]. Despite the initial induction of p53 in nickel-exposed cells, the functional activity of p53 is decreased in nickel-transformed cells. Additionally, in human and rodent nickel-transformed cells, a shift in the balance toward HIF-1-dependent transcription and weakening p53-dependent transcription was observed [125].

6.7.

Retinoblastoma

Rb is the first tumor suppressor gene identified through human genetic studies. This nuclear protein was originally found to be lost or mutated in retinoblastomas [197]. Since Rb serves as a transducer between the cell cycle machinery and promoter-specific transcription factors, Rb mutation may result in deregulation of the cell cycle and subsequent cell proliferation. A substantial number of the Rb-interacting proteins are transcription factors such as E2F, Elf-1, DRTF-1, and NF-IL6 [197]. In general, these transcription factors are inactive when they are bound to a hypophosphorylated form of Rb. Once Rb is phosphorylated, transcription factors are released and ready to interact with other transcription factors and eventually with DNA to stimulate gene expression. Thus, two types of modifications of Rb are important in carcinogenesis; first, modification or loss of the protein due to mutation or deletion; second, abnormal phosphorylation of Rb protein. Nickel-induced retinoblastomas were first described when nickel subsulfide, Ni3S2, was administered to albino Fischer rats by a single injection into the vitreous body of the eye [198]. The status of the Rb protein was not tested in those experiments. In the in vitro model, when human osteosarcoma (HOS) cells, which do not grow in soft agar or form tumors in athymic nude mice, were repeatedly treated with water-soluble NiSO4, or water-insoluble NiS, an increase in anchorage-independent colony formation was observed [21,196]. In eight of nine nickel-transformed clones obtained, Rb protein was found in the hypophosphorylated form and mutational changes of the Rb gene in some of the nickeltransformed clones were identified [196]. It is not known, however, whether these mutations are the result of nickel exposure or are selected in nickel-transformed cells as the result of soft-agar selection. When the normal Rb phosphorylation pattern was restored by the transfection of the plasmid containing the wild-type Rb gene, the cells acquired a normal phenotype (i.e., they were no longer able to grow in soft agar). The mechanism for the abnormal Rb phosphorylation (one would expect to find hyperphosphorylated forms in nickel-transformed cells) is not known. Neither is the nickel specificity of this effect. Additionally, it was also found that the level of Rb expression in nickel-transformed HOS clones was decreased. Met. Ions Life Sci. 2, 581–618 (2007)


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6.8. FHIT The FHIT (for fragile histidine triad) gene is a tumor suppressor gene located in a fragile chromosomal site sensitive to deletions. Therefore, its expression is frequently reduced or lost in tumors and pre-malignant lesions. Silencing of the FHIT tumor suppressor gene through DNA methylation is one of the inactivation mechanisms [199]. Its product, FHIT protein (phosphohydrolase), induces apoptosis through complex interactions with its substrate, diadenosine triphosphate (Ap3A) [200]. Ni(II) was found to strongly inhibit the enzymatic activity of FHIT protein in vitro and also suppress FHIT’s expression in nickel-transformed BALB/c-3T3 cells [201]. In these cells, FHIT protein levels were reduced by 50%, compared with those in the parental cells. A decrease in FHIT protein levels by up to 90% was also observed in 22 local sarcomas induced by intramuscular injection of Ni3S2 in mice, as compared with normal mouse muscles [202]. Moreover, FHIT was absent in 4 of those sarcomas. The decrease in FHIT expression coincided with faster development of tumors. Overall, the decline of FHIT in cells or tissues malignantly transformed by nickel, along with inhibition by nickel of FHIT’s enzymatic activity, may indicate a possible contribution of these two effects to the mechanisms of nickel carcinogenesis [202].

7.

CHANGES IN GENE EXPRESSION AND NICKEL CARCINOGENESIS

The use of microarrays in analyses of gene expression in nickel-exposed cells has increased over the past few years, providing expression profiles for a large number of genes in a short time [24,57,64,87,128,129]. Transgenic and knockout mouse models made it possible to study the in vivo function of genes regulated by nickel exposure and their products. These techniques revealed a multitude of causes for aberrant gene regulation following acute nickel exposure or in nickel carcinogenesis. Gene deregulation can occur at different levels – the genome level, the level of signaling and transcription and even the post-translational level. One of the alterations occurring at genome level is cytogenetic changes. These changes are noticeable for the heterochromatic long arm of the X chromosome and are described above in Section 2. Another mechanism of gene inactivation in nickel carcinogenesis is epigenetic silencing by DNA methylation. In mammals, DNA methylation is predominantly observed in CpG dinucleotides that occur at a higher frequency than expected in stretches of DNA known as CpG islands. The latter are commonly found in promoters or other cis-regulatory regions. At these locations, methyltransferases can methylate the cytidine residues, while under normal conditions, methylation of these cytidines is restricted to particular physiologic situations such as embryogenesis Met. Ions Life Sci. 2, 581–618 (2007)

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and cell differentiation. These changes can be induced by nickel and are described in Section 2.1. In spite of identification of these changes in some model systems, it is unclear whether a change in the gene methylation status is a general phenomenon of nickel exposure or a gene-specific phenomenon. In the case of hypermethylation observed in vivo, it is not clear whether hypermethylation is a cause or a consequence of the malignant phenotype produced by nickel exposure. The ‘histone code’ is a widely accepted hypothesis, whereby sequential modifications to the histones in chromatin lead to regulated transcription of genes. One of the modifications used in the histone code is acetylation. This is probably the best characterized modification of histones, which is carried out under the control of histone acetyltransferases (HATs) and histone deacetylases (HDACs). These enzymes also regulate the activity of a number of transcription factors through acetylation. Increasing evidence links nickel-induced dysregulation in gene expression and inhibition of histone acetylation [203]. These mechanisms possibly play an important role in the pathogenesis of nickel-induced diseases. Finally, altered signaling pathways or deregulated transcription factors represent an important category of molecular events leading to aberrant gene regulation in nickel carcinogenesis. Nickel compounds have profound effect on complicated cellular signaling pathways, which results in post-translational modifications of numerous transcription factors. Understanding how the overall changes in cellular signaling network translates into activation or suppression of transcription factors and regulation of specific subsets of genes will lead to a mechanistic understanding of how nickel mediates its diverse and paradoxical biological effects.

8.

CONCLUSIONS

Multiple signaling pathways are induced following acute nickel exposure. They may turn on both protective and damaging mechanisms. Depending on the dose of nickel, which determines the extent of injury, one of these mechanisms prevails. A significant role in nickel-induced in vivo acute toxicity plays the induction of cytokines, which amplify the response in case of lung injury or allergic dermatitis. During chronic exposure, in addition to the induction of inflammatory response, there is an induction of genes coding for proteins involved in hypoxic stress that shifts cellular metabolism to a glycolytic pathway. DNA damage and the inhibition of DNA repair, may synergistically favor neoplastic growth of nickel-exposed cells. Microarrays as molecular tools are currently used to profi le changes in cells exposed to nickel compounds and to produce ‘molecular signatures’. These signatures may reflect genetically and epigenetically mediated changes in patterns of gene expression, and provide new opportunities for identification of relevant biomarkers of nickel exposure, which have potential for monitoring Met. Ions Life Sci. 2, 581–618 (2007)


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the levels of exposure of workers in nickel-related industries. They are also instrumental in the development of new molecular diagnostic and treatment strategies.

ACKNOWLEDGMENTS This work was supported by the Intramural Research Program of the NIH, NCI.

ABBREVIATIONS AhR Aqp1 ARNT BAL BAPTA-AM CCL CHO COX-1 and COX-2 C-P4H DCFH DMOG DMT-1 ECM ELAM-1 Epo EST FHIT FIH GAPDH GSH GST IARC ICAM-1 IL HAT HDAC HIF HO-1 HOS HRE

aryl hydrocarbon receptor aquaporin-1 aryl hydrocarbon receptor nuclear translocator bronchoalveolar lavage 1,2-bis-(o-aminophenoxy)-ethane-N,N,N′,N′-tetraacetic acid, tetraacetoxymethyl ester chemokines Chinese hamster ovary cells cyclooxygenase 1 and 2 collagen prolyl-4-hydrolase 2′,7′-dichlorofluorescin dimethyloxalylglycine divalent metal transporter 1 extracellular matrix endothelial leukocyte adhesion molecule-1 erythropoietin expressed sequence tag fragile histidine triad factor inhibiting HIF glyceraldehyde-3-phosphate dehydrogenase glutathione glutathione-S-transferase International Agency for Research on Cancer intercellular adhesion molecule-1 interleukins histone acetyltransferase histone deacetylase hypoxia-inducible factor heme oxygenase-1 human osteosarcoma cells HIF response element Met. Ions Life Sci. 2, 581–618 (2007)

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MCP-1 MT-1 ODD 2OG PAI-1 PBMC PI3K PMA Rb ROS RT-PCR SCE SP SVCT2 TGF TNF-α TSA TSP I VCAM-1 VEGF VHL VLA-3


monocyte chemoattractant protein-1 metallothionein-1 oxygen-dependent degradation domain 2-oxoglutarate plasminogen activator inhibitor peripheral blood mononuclear cell phosphatidylinositol-3-kinase phorbol ester ⫽ phorbol 12-myristate 13-acetate retinoblastoma reactive oxygen species real-time polymerase chain reaction sister chromatid exchange surfactant proteins sodium-dependent vitamin C transporter transforming growth factor tumor necrosis factor α trichostatin A thrombospondin I vascular cell adhesion molecule-1 vascular endothelial growth factor von Hippel Lindau very late antigen-3

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17 Nickel Toxicity and Carcinogenesis Kazimierz S. Kasprzak and Konstantin Salnikow Laboratory of Comparative Carcinogenesis, National Cancer Institute at Frederick, Frederick, MD 21702, USA <salnikow@ mail.ncifcrf.gov >

1. INTRODUCTION 2. AN OVERVIEW OF NICKEL TOXICITY 2.1. Toxicity of Nickel in Humans and Domestic Animals 2.1.1. Routes of Exposure, Uptake, and Excretion 2.1.2. Acute Exposures 2.1.3. Chronic Exposures 2.1.3.1. Non-Malignant Respiratory Lesions 2.1.3.2. Nickel Dermatitis 2.1.4. Detoxification 2.1.5. Nickel Deficiency 2.2. Toxicity of Nickel in Experimental Animals 2.2.1. Uptake, Distribution, Metabolism, and Excretion 2.2.2. Systemic Toxicity 2.2.3. Organ and Tissue Toxicity 2.2.3.1. The Respiratory Tract 2.2.3.2. Liver 2.2.3.3. Kidneys 2.2.3.4. Cardiovascular System 2.2.3.5. Nervous System 2.2.3.6. Immune System 2.2.3.7. Reproductive System and Embryotoxicity 2.3. Molecular Mechanisms of Nickel Toxicity Metal Ions in Life Sciences, Volume 2 © 2007 John Wiley & Sons, Ltd


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Binding to Biomolecules 2.3.1.1. Nickel Carriers 2.3.1.2. Regulatory and Structural Proteins 2.3.1.3. Histones and Protamines 2.3.2. Mediation of Oxidative Damage 2.3.2.1. Lipid Peroxidation 2.3.2.2. Protein Damage 2.3.2.3. DNA Damage 2.3.2.4. Effect on Ascorbic Acid, Glutathione, and Other Antioxidants 3. NICKEL-INDUCED CARCINOGENESIS 3.1. Human Epidemiology 3.1.1. Carcinogenic Nickel Species 3.1.2. Histopathology of Nickel Cancer 3.2. Carcinogenesis in Experimental Animals 3.2.1. Carcinogenic Nickel Compounds 3.2.2. Administration Routes 3.2.2.1. Intramuscular and Subcutaneous Injections 3.2.2.2. Intraperitoneal Injection 3.2.2.3. Intrarenal Injection 3.2.2.4. Intratesticular Injection 3.2.2.5. Intraocular Injection 3.2.2.6. Inhalation 3.2.2.7. Other Routes of Exposure 3.2.3. Species and Strain Susceptibility 3.2.4. Interactions with Other Carcinogens 3.2.5. Interactions with Essential Metals 3.3. Malignant Transformation of Cultured Cells 3.4. Mechanisms of Nickel-Induced Carcinogenesis 3.4.1. Genotoxic Effects 3.4.2. Epigenetic Effects 3.4.3. Molecular Basis for the Genetic and Epigenetic Toxicity of Nickel 3.4.3.1. Promutagenic Effects of Oxidative DNA Damage 3.4.3.2. Inhibition of DNA Repair Enzymes 3.4.3.3. Effects on DNA Methylation and Histone Modifications 3.4.3.4. Disruption of Calcium Homeostasis 4. CONCLUSION ACKNOWLEDGMENTS ABBREVIATIONS REFERENCES Met. Ions Life Sci. 2, 619–660 (2007)

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NICKEL TOXICITY AND CARCINOGENESIS

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While nickel is essential in certain bacteria, plants, and domestic animals, its essentiality in humans remains doubtful. Nickel toxicity reports (in ore miners) date from the 1500s, i.e., even before elemental nickel was first identified [1]. The first experimental studies of nickel toxicity in animals are ascribed to Gmelin (1826) and Stuart (1884) [2]. Clinical, epidemiological, and experimental investigations of nickel toxicity increased rapidly after nickel carbonyl was discovered (Mond, 1889) and used in nickel metallurgy, exposing hundreds of workers to this highly toxic volatile liquid. The death or acute respiratory distress following nickel carbonyl inhalation, observed at the beginning of its use, were soon prevented by technical measures; however, the adverse effects, such as ‘nickel dermatitis’ and respiratory tract cancer from chronic low-dose exposures continue as health problems today [3]. To resolve these health issues, full recognition of all symptoms and understanding of the mechanisms of nickel toxicity are necessary, and require the help of experimental models, including animals, cultured cells, cell-free systems, and computational techniques, as reviewed below.

2.

AN OVERVIEW OF NICKEL TOXICITY

2.1.

Toxicity of Nickel in Humans and Domestic Animals

Nickel toxicity in humans has been reviewed before [4–13]. Since nickel is widely used in modern industries, the exposures and associated health problems are predominantly occupational, but contact with domestic or personal products containing nickel, e.g., jewelry, or internal surgical and dental devices may result in allergic or inflammatory reactions [14,15], depending on the compound, dose, administration route, and exposure time.

2.1.1.

Routes of Exposure, Uptake, and Excretion

In nickel metallurgy and related industries the major route of nickel entry into the body is the respiratory tract. Oral ingestion and dermal absorption are of secondary importance; they do, however, become the only significant routes for general environmental exposures [4–13]. There are also reports of iatrogenic exposures [16]. Studies of nickel absorption and elimination in workers inhaling nickelcontaining dusts and aerosols have been reviewed by Sunderman et al. [17,18]. Nickel inhaled as a water-soluble compound is absorbed into the blood that transports it to other tissues, and excreted in urine, with T1/2 of 17–39 h [19]. Following inhalation of water-insoluble nickel dusts and fumes, nickel is eventually dissolved in tissue fluids, mobilized by blood, and eliminated in urine with T1/2 of Met. Ions Life Sci. 2, 621–662 (2007)

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30–53 h [18]. These data might imply fast elimination of inhaled nickel; however, low amounts of nickel can persist for months and years after exposure in tissue ‘sinks’, as found in retired nickel workers’ nasal epithelia [18]. Animal experiments show that lung and kidneys may also constitute such sinks [20–22]. Inhalation of the volatile nickel carbonyl is especially dangerous because of its extremely high toxicity [5]. This nonionic lipid-soluble nickel compound easily crosses cell membranes, is metabolized by oxidation to water-soluble Ni(II) and CO ⫹ CO2, and eventually excreted in urine [23,24]. Larger nickel particles may also be deposited in the nasal cavities and then mobilized into the body. Sunderman [25] found that nickel can be transported from the nose via olfactory axons to the olfactory bulb of the brain. Studies of gastrointestinal absorption of water-soluble nickel in humans revealed a relatively fast absorption into the blood, peaking around 2.5–3 h post-ingestion, and returning to normal level in approximately 72 h (T1/2 ⬃ 11 h). Because of the sinks and other factors (e.g., water intake and diuresis), the bodily half-time of nickel ingested in a water-soluble form may vary widely, from 11 to over 30 h. Food taken with nickel greatly limits its absorption [26]. In persons who fasted overnight before taking soluble nickel, the absorption ranged from 29 to 40% of the dose; urinary excretion amounted to 51–82% in 5 days; plasma nickel peaked between 1.5 and 2.5 h [27]. There is no published data on the absorption of orally taken water-insoluble nickel compounds in humans. In the bloodstream, plasma proteins (mainly albumin) and poorly defined lowmolecular-mass ligands carry nickel throughout the body [28]. Bones and lungs tend to retain it relatively longer than do other tissues. Kidney is the major excretory organ and, therefore, the primary subject of absorbed nickel toxicity. Unfortunately, little is known about the specific distribution of toxic doses of nickel in human tissues as compared with that in experimental animals [29] (compare Section 2.2.1). Skin exposure may result in nickel absorption through sweat ducts and hair follicles. Sweat and detergents may speed this process up by enhancing the dissolution of nickel from water-insoluble dusts or solids, and increasing skin permeability. Most of the absorbed nickel remains bound to keratin in the dermis [5]. Symptoms of nickel poisoning were observed in patients undergoing hemodialysis. The source of nickel was contaminated water. Intravenous infusion of nickel-contaminated medications and corrosion of internal prostheses can also deliver toxic nickel doses or trigger nickel allergies [16,30].

2.1.2.

Acute Exposures

Except that of nickel carbonyl (see below), the toxicity of nickel compounds is not high; therefore acute poisoning is rare [31,32]. Oral doses of water-soluble nickel salts below 1 g Ni may induce nausea, slow heart pulse, and decrease body Met. Ions Life Sci. 2, 619–660 (2007)


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temperature. Higher doses, causing hemorrhagic gastritis, pulmonary congestion, tachycardia, and eventually cardiac arrest, may be fatal [33]. Sunderman et al. [32] reported an accident in an electroplating shop involving 32 workers who drank water contaminated with nickel sulfate and chloride. Twenty promptly developed nausea, vomiting, abdominal pain, diarrhea, dizziness, weakness, headache, cough, and shortness of breath that persisted for up to 2 days; nickel doses ranged from 0.5 to 2.5 g Ni per person. All victims recovered within eight days without any evident sequels. The extremely high toxicity of nickel carbonyl is unique among nickel compounds. It exceeds the toxicity of carbon monoxide by a factor of 100 [5] and is comparable to that of hydrogen cyanide. Atmospheric concentration of nickel carbonyl vapor of 30 ppm is lethal for man after a 30-min exposure [34]. The initial effects include irritation of the respiratory and gastrointestinal tracts and central nervous system. Severe delayed symptoms such as breathing problems, tachycardia, muscular pain and weakness, abdominal pain, and diarrhea, may occur 12–36 h after exposure [5,35–37]. Pneumonitis and cerebral edema develop in severe cases and may cause death [38]. In men who died 4–13 days after the exposure, brain hemorrhages, edema and interstitial fibrosis of the lung, and in some cases also degeneration of liver, kidneys, adrenal glands, and spleen, were observed [5].

2.1.3.

Chronic Exposures

Prolonged exposures to low doses of nickel may produce severe pathologic effects, including dermatitis, asthma, lung fibrosis, and respiratory tract cancer [5,12,31,38,39]. Although generally nickel has been regarded as a noncumulative toxin in humans and animals, some adverse changes in renal function observed in nickel workers [40] showed dose–response trends typical for cumulative exposures [41]. 2.1.3.1. Non-Malignant Respiratory Lesions. Inhalation of nickel carbonyl vapors or nickel-containing aerosols may lead to respiratory insufficiency and lung fibrosis [16]. Berge and Skyberg [41] identified lung fibrosis in 47 of 1046 workers handling various nickel compounds in a nickel refinery. Significant risk of developing the fibrosis was found for workers exposed to both insoluble (sulfidic) and water-soluble nickel, but not for metallic nickel. There was a clear dose–response trend for soluble, but not for insoluble nickel. Asthma and other chronic respiratory ailments such as bronchitis, sinusitis and rhinitis (anosmia), have also been observed among nickel refinery workers [25,42,43]. The asthmatic reaction to nickel develops independently from the more frequent dermatitis. However, both may share a common primary antigen: Ni(II) bound to serum albumin [44]. Nickel also induces metal fume fever [43,45] and Loffler’s syndrome [46,47]. Met. Ions Life Sci. 2, 621–662 (2007)

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2.1.3.2. Nickel Dermatitis. Asthma, metal fume fever, and Loffler’s syndrome indicate the capacity of nickel to attack the immune system. Other examples include conjunctivitis, stomatitis, and inflammatory reactions around implants or orthodontic appliances made of nickel alloys [14,15,48]. However, the most prevalent result is allergic contact dermatitis, making nickel one of the most potent human allergens [5,49]. Although occupational skin exposures are mainly to blame, sensitization to nickel is frequently due to non-occupational encounters with nickel-containing jewelry, clothing fasteners, bracelets, wristwatches, dental prostheses, and internal surgical devices; women are more sensitive than men [5,10]. Symptoms may persist for months after contact with nickel, a possible result of long-term nickel sequestration by keratin [50].

2.1.4.

Detoxification

Acute nickel toxicity symptoms can be alleviated by chelation therapy. For example, intramuscular administration of 2,3-dimercaptopropanol to workers exposed to nickel carbonyl resulted in increased nickel excretion in urine and some clinical benefits [51]. Penicillamine gave equivocal results [52,53]. Thus far, the most effective antidote in clinical treatment of nickel carbonyl poisoning appears to be sodium diethyldithiocarbamate (dithiocarb) introduced by Sunderman and Sunderman in 1958 and successfully administered to hundreds of patients (reviewed in [54]). Disulfiram, a metabolite of dithiocarb, has also been used [55]. However, a therapeutic review of these chelators indicates that to be effective dithiocarb must be administered parenterally soon after exposure to nickel carbonyl. It also points at the alarming fact that high doses of disulfiram increased mortality of experimental mice given nickel carbonyl, most likely by enhancing nickel accumulation in the brain [56]. The latter conclusion is consistent with animal and in vitro observations of Nieboer et al. [57,58]. Thus, more studies are required before dithiocarb can be recommended for a routine clinical use. Diethylenetriaminepentaacetic acid (DTPA) has been used successfully in patients to prevent allergic contact dermatitis caused by nickel, cobalt, or copper [59]. Numerous other chelators were tested for their efficacy in removing nickel from the body, but only in animals; no definitive relationship between their structure and effectiveness was found [60,61]. Blanusa et al. recently reviewed the use of chelators against metal toxicity [62].

2.1.5.

Nickel Deficiency

The role of the nickel-bearing protein, nickeloplasmin, in human blood [63] remains unknown. Thanks to the dietary and environmental exposures, low amounts of nickel are found in virtually all human tissues; therefore, nickel deficiency, if any, can hardly occur [64]. In contrast, nickel essentiality has been Met. Ions Life Sci. 2, 619–660 (2007)


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clearly established in animals (reviewed in [65]). Rats fed a nickel-free diet exhibited lower reproductive performance, increased perinatal mortality, impaired liver function, growth retardation, and reduced life span. The second generation of these rats had anemia due to impaired dietary iron absorption; the anemia worsened significantly in the third generation [66, 67]. Goats fed a nickel-deficient diet showed growth retardation, increased mortality of dams and offspring, parakeratosis of the skin; lower activities of several key enzymes in the liver, kidneys, and especially heart; and degeneration of cardiac and skeletal muscles [68]. More recently, Yokoi et al. [67] found that the detrimental effects of nickel deficiency might be due to impaired function of cyclic nucleotide-gated (CNG) ion channels.

2.2. Toxicity of Nickel in Experimental Animals The symptoms of nickel toxicity in animals following oral, inhalatory, and dermal exposures are virtually the same as in humans. The differences in toxic activities observed among nickel compounds depend on the mode of exposure, dose, uptake, transport, distribution, and retention, and – ultimately – on the capacity of a given compound to interact with specific cellular and molecular targets. These factors, in turn, depend on the solubility, particle structure and size, and redox activity of different nickel forms. Using experimental animals enables testing of the importance of all those factors. The major advantage of animal studies stems from the option of parenteral nickel administration.

2.2.1.

Uptake, Distribution, Metabolism, and Excretion

The pharmacokinetics of nickel at the whole-body and tissue levels have been studied experimentally in animals and tested in mathematical models [69,70]. In studies, water- and acid-soluble nickel salts ingested orally were excreted mainly in the feces, with only 1.7–10% of the dose excreted in urine. At 45–75 h after oral ingestion of soluble nickel, its whole-body retention in mice was only 0.02–0.36% of the dose. At 8 h, the highest concentration of the absorbed nickel was in the kidneys, followed in decreasing levels by the carcass ⬎ lungs ⬎ testes ⬎ liver ⬎ spleen. At 20 h, nickel was present in the carcass ⬎ kidneys ⬎ liver ⬎ lungs. At 40 h, the hepatic nickel level exceeded that of renal nickel [71]. After a single oral dose of soluble nickel to mouse, Borg and Tjalve [72] found low levels of nickel in the sciatic nerves, trigeminal ganglia, spinal nerve roots, spinal cord, cerebellum, and frontal cortex of the brain. Prolonged exposure of rats to nickel in drinking water increased nickel levels in tissues and body fluids; the highest level was found in the liver, followed by kidney ⫽ blood ⬎ serum ⬎ testes ⬎ urine ⬎ ovaries [73]. Oral ingestion of insoluble nickel compounds resulted in much lower intestinal absorption [74]. Met. Ions Life Sci. 2, 621–662 (2007)

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Inhalation of nickel carbonyl by animals increases the lung and body burden of nickel that is subsequently excreted, mainly in urine [23,75]. Likewise absorbed from the lungs into the bloodstream is nickel from inhaled water-soluble and insoluble nickel aerosols [76–78]. However, the lung clearance is slow, with 28% of the initial dose still present in the lung four days after inhalation of nickel chloride [76]. Inhalation of insoluble nickel compounds results in longer retention and multi-phasic mobilization of nickel from the lung with half-lives ranging from several days to several months, depending on the dose and nature of the particles [79–81]. Nickel from salts painted on the skin is mainly sequestered in the dermis, but hepatic and testicular lesions found in rats after dermal exposure indicate the possibility of systemic distribution [82]. Nickel also was found to cross the placental barrier, accumulate in the embryos or fetuses, and increase lethality and malformations of the offspring [83,84]. Administering nickel compounds intravenously, intraperitoneally, subcutaneously, or intramuscularly results in the blood absorbing nickel quickly and redistributing it among various organs and tissues. Sunderman and Selin [23] found that most of the nickel chloride injected intravenously in rats was quickly excreted in urine: 87% in one day, 90% in two days, plus about 3% in feces. Further excretion was much slower, indicative of long-term nickel retention in certain tissues. A single intravenous or intraperitoneal injection of 63Ni(II) chloride to mice allowed for nickel distribution and retention in tissues of mice to be traced using whole-body autoradiography. In the first few hours after injection, nickel was taken up from blood by connective tissues and cartilages and was also present in high amounts in the kidneys and urinary bladder. One day after the injection, nickel radioactivity decreased in blood and skin, but remained high in the kidney and cartilages; at the same time, radioactivity in the forestomach and, especially, in the lungs markedly increased. Five days later, nickel was still present in the lungs, kidneys and forestomach, although at levels lower than in the spinal cord and brain. Micro-autoradiography of the lung revealed an accumulation of 63Ni mainly in the lung parenchyma and lower amounts in the bronchial epithelium. Nickel in the forestomach was sequestered by the keratinized squamous epithelium. In the kidney, 63Ni was localized in small areas of the cortex and distal convoluted tubules, and also in the pelvis and in the zone between the medulla and cortex. Oral exposure resulted in high 63Ni contents in the gastrointestinal tract, with only a small part of the dose being absorbed into the bloodstream and distributed in the same way as after parenteral injections [20]. The 63Ni and 14C labels in nickel carbonyl revealed that this compound could not only enter, but also leave the animal body through the lung [23,24]. Over 35% of nickel carbonyl dose injected intravenously to rats was exhaled within the first 4 h. The rest dissociated and was oxidized to Ni(II), CO, and CO2. In the first 6 h post injection, approximately 50% of the carbonyl moiety was exhaled as CO Met. Ions Life Sci. 2, 619–660 (2007)


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and 1% as CO2. At the same time, most of the nickel moiety was retained in the lung, followed by kidneys, urinary bladder, brain and spinal cord, heart muscle, diaphragm, brown fat, adrenal cortex, and ovaries. This indicated that Ni(II) derived from nickel carbonyl followed the distribution and excretion pathways of other soluble nickel compounds [22]. 63Ni- and 35S- also allowed for tracing the uptake and metabolism of carcinogenic nickel subsulfide, Ni3S2, injected intramuscularly in mice or rats [85]. The injected Ni3S2 particles slowly lost radioactivity of both markers due to solubilization, phagocytosis, and translocation to regional lymph nodes, spleen, and liver. The particles remained detectable in tumors, which developed at the injection sites after more than 6 months. X-ray crystallography revealed conversion of Ni3S2 to Ni7S6 and NiS, that was consistent with Ni3S2 oxidation and solubilization reactions observed before in vitro [86]. According to Hildebrand et al. [87], the sulfur moiety of intracellular Ni3S2 is gradually replaced by phosphorus of different organic phosphates. Studies on cellular and subcellular uptake and retention of nickel revealed that Ni2⫹ cations may diffuse through cell membranes [88] or be actively transported via calcium [89] and iron channels [90]. The latter is likely to involve a proton-coupled divalent cation transporter DMT-1 (Nramp 2) that may account for the observed transport antagonism between nickel and other cations, including iron [91]. Ni2⫹ interactions with iron transport and storage also seem possible at the transferrin/ferritin system [92]. Overall, these ways of nickel uptake, e.g., in the gastrointestinal tract, are inefficient and concur with low toxicity of water-soluble nickel compounds. The third mechanism of cellular nickel uptake involves phagocytosis of nickel dusts, observed in cultured cells; its efficiency depends on the size and electric charge of the particles [93,94]. Crystalline Ni3S2, NiS, and Ni3Se2 particles smaller than 5 µm, engulfed by cells, formed soluble Ni2⫹-generating vacuoles localized close to the nucleus [95]. These findings are consistent with studies showing that nickel released from Ni3S2 and NiO particles reaches the nucleus in greater amounts than nickel from water-soluble Ni(II) sulfate [9,96]. For toxicity to occur it is essential, however, that such particles are dissolvable intracellularly, as is the case of Ni3S2 and NiS [85,86,97]. Measurements of nickel in subcellular fractions showed that exposure of cells to water-soluble salts produced relatively high cytosolic, but very low nuclear nickel levels, while exposure to crystalline Ni3S2 resulted in high nickel contents in both cytosolic and nuclear fractions [96]. In the lung of mice given parenteral injections of nickel acetate, the highest nickel levels were found in the microsomes ⬎ mitochondria ⬎ cytosol ⬎ nuclei [98].

2.2.2.

Systemic Toxicity

Orally ingested nickel is acutely toxic only in relatively high doses, depending on the species. For example, LD50 for nickel acetate is 350 mg/kg body weight Met. Ions Life Sci. 2, 621–662 (2007)

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in rats and 420 mg/kg body weight in mice [4,5]. Extensive data on the toxicity of nickel compounds may be found in [5]. Inhalation of nickel carbonyl is as lethal in animals as in humans: LD50 for a 30-min exposure was 0.067, 0.24, or 0.19 mg/L of air for mouse, rat, or cat, respectively. Death was also observed in rats and mice following inhalation of high doses of nickel sulfate (⬎30 mg Ni/m 3 for less than 12 days) [99]. Parenteral administration makes nickel much more toxic and invokes some specific effects in individual organs and tissues. For example, the fi rst consequence of parenteral nickel injection is acute hyperglycemia that is most likely due to glucagon release from the pancreas [100].

2.2.3.

Organ and Tissue Toxicity

2.2.3.1. The Respiratory Tract. Rats and mice that inhaled water-soluble nickel sulfate aerosols developed lung inflammation, hyperplasia of bronchial and mediastinal lymph nodes, and atrophy of the olfactory epithelium, which worsened with increase in dose and time of exposure. Alveolar proteinosis and lung fibrosis developed in animals inhaling nickel for 13 weeks or longer [99]. Inhalation of aerosols of the insoluble nickel oxide and nickel subsulfide produced pathologic effects very similar, although of greater severity, to those induced by the soluble nickel sulfate; also, mice were somewhat less sensitive than rats. However, unlike nickel sulfate, these compounds were also carcinogenic to the lung [101,102]. Lung inflammation induced by nickel aerosols was also observed in monkeys, rabbits, and hamsters [103]. Since lung is a major excretory organ for nickel carbonyl, it is also the primary target for the toxicity of this compound, regardless of the administration route. The immediate symptoms include edema and leukocytic infiltration of the peribronchial and alveolar septal interstitium, followed by proliferation and hyperplasia of bronchiolar epithelium and alveolar lining cells. During the next 4 days, severe hemorrhagic intra-alveolar edema may develop and lead to death of the animal. In surviving animals, the cytologic changes regress up to a full recovery (except for interstitial fibrosis) in 2–3 weeks after exposure [5]. 2.2.3.2. Liver. In rats, subcutaneous administration of high doses of nickel chloride resulted in acute hepatic toxicity with fatty metamorphosis, hydropic degeneration, and focal inflammation developing in 1–3 days. Lipid peroxidation and activities of serum aminotransferases, indicative of liver damage, were greatly enhanced [104]. Multiple intramuscular injections of low doses of nickel sulfate caused similar liver damage [105]. In mice, after intraperitoneal injection of a single dose of nickel acetate, the extent of lipid peroxidation depended on the decrease by nickel of the glutathione and glutathione peroxidase and reductase levels in the respective strain [106]. Met. Ions Life Sci. 2, 619–660 (2007)


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2.2.3.3. Kidneys. Aminoaciduria and proteinuria developed promptly in rats given a single intraperitoneal injection of nickel chloride; the symptoms subsided in 5 days [107]. Similar symptoms were observed shortly after inhalation of nickel carbonyl [108]. A chronic exposure of rats to 1.7 mM nickel sulfate in drinking water for 6 months caused kidney enlargement and proteinuria. Female rats were more sensitive to these effects than were male rats. Kidney damage was also observed in nickel chloride-injected rabbits [109]. The functional damage of kidney by nickel, resulting in proteinuria, has been associated with nickel binding to the glomerular basement membrane [110]. A unique effect of nickel on the kidney is the long-lasting erythrocytosis following intrarenal injection of Ni3S2 and some other insoluble nickel derivatives in rats. This reaction, due to induction of erythropoietin, involves erythroid hyperplasia of bone marrow and spleen [111–114]. Guinea pigs, but not hamsters and gerbils, respond to intrarenal nickel subsulfide in the same way [115]. Morphological effects of Ni3S2 in the kidney include glomerulomegaly, tubular hyperplasia, and arteriosclerosis [112]. 2.2.3.4. Cardiovascular System. Nickel reduced the contractility of cardiac muscle in rat, guinea pig, and dog, and in isolated canine coronary artery (reviewed in [10]), probably because of nickel’s interaction with calcium and sodium channels [116]. Elevated nickel released to the bloodstream from damaged tissues, e.g., in burns, as well as nickel contaminating infusion fluids, may therefore induce vasoconstriction and myocardial injury [117,118]. Intrarenal injection of nickel subsulfide in rats caused widespread arteriosclerotic lesions 8–18 weeks later. Arteriosclerotic plaques were found in all major arteries despite the lack of hypertension and hyperlipidemia as pathogenic factors [119]. 2.2.3.5. Nervous System. Rats subject to chronic inhalation of nickel sulfate aerosol showed atrophy of olfactory epithelium with reduced numbers of bipolar sensory neurons and levels of the neurochemical marker carnosine, but without measurable changes in olfactory function [120]. Repeated daily intraperitoneal injections of soluble nickel to rats resulted in cerebral edema and increased lipid peroxidation that was highest in the cerebellum, followed by brain stem and the hemispheres [121]. 2.2.3.6. Immune System. Sensitization of animals to nickel through dermal exposure (topical or intradermal) produced positive results in guinea pigs and mice. Inflammation of skin caused by other insults appeared to increase the response to nickel [5,122–125]. Artik et al. [126] found that nickel at higher oxidation states, Ni(III) or Ni(IV), was more allergenic than Ni(II). This may explain why nickel dermatitis develops more readily in inflamed skin, i.e., under Met. Ions Life Sci. 2, 621–662 (2007)

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conditions favoring Ni(II) oxidation, than in normal skin. The development of allergic dermatitis involves presentation of the nickel antigen by Langerhans cells and its recognition by T-lymphocytes [10,127]. In vitro stimulation by nickel of guinea pig lymphocytes and thymocytes has been reported [128]. Nickel may also attack other types of cells of the immune system. For example, nickel’s effect on alveolar macrophages, avidly scavenging nickel-containing particles, was compared experimentally in mice, rats, rabbits, and dogs [10]. Up to a certain level of exposure the macrophages are stimulated and their size, number, and phagocytic activity increase, but higher exposures are deadly. An intramuscular injection of nickel chloride in mice caused an involution of the thymus, ‘chemical thymectomy’. In the spleen, the number of T-lymphocytes decreased, while that of B-lymphocytes remained unchanged. Macrophage function was not markedly affected, but the cytotoxic activity of natural killer (NK) cells was diminished [129,130]. The acute thymic involution [131] and suppression of NK cells was also observed in rats given parenteral injections of nickel chloride and nickel subsulfide [132,133] or intratracheal instillation of nickel sulfate [134]. NK cells’ suppression by nickel was also observed in monkey cells in vitro [135]. However, Ni3S2 instilled in a monkey lung activated NK cells [136]. Nickel was also found to increase the pathogenicity of Coxsackie virus infecting murine myocardium, lung, and pancreas through suppression of cytotoxic T cells, helper T cells, and macrophages [137].

2.2.3.7. Reproductive System and Embryotoxicity. Adding nickel sulfate to drinking water for mice and rats reducted fertility, decreased litter size, and increased offspring mortality [138]. Morphologic sperm abnormalities, reduced sperm motility and sperm count, and alterations in the activities of marker testicular enzymes were also observed [139]. Nickel may halt sperm motility through a selective inhibitory effect on dynein [140]. Subcutaneous injections of nickel sulfate to rats caused hyperemia of testicular intertubullar capillaries, disintegration of spermatozoa, and inhibition of spermatogenesis, depending on the dose [141,142]. Domshlak et al. [143] found that in mice spermatozoids, early spermatids, late spermatocytes, and stem spermatogonia are most sensitive to nickel sulfate. On the other hand, nickel deprivation also reduced the rat reproductive performance by decreasing sperm production rate and spermatozoa motility. Nickel deficiency also significantly reduced the weights of the seminal vesicles and prostate glands [67]. Since nickel crosses the placental barrier, it may harm the embryo or fetus. Experiments in mice, rats, and hamsters revealed that maternal exposure to nickel decreased implantation frequency, caused early and late resorptions, and increased the number of stillborn and abnormal offspring. The teratogenic effects of nickel included acephalia, exencephaly, cerebral hernia, skeletal anomalies, micromelia, microphthalmia, anophthalmia, and others [10, 144]. Microphthalmia and Met. Ions Life Sci. 2, 619–660 (2007)


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anophthalmia were also observed in chickens and tadpoles exposed to nickel during embryogenesis [145,146].

2.3. Molecular Mechanisms of Nickel Toxicity 2.3.1.

Binding to Biomolecules

At the physiological pH range, the strength of nickel Ni2⫹ interactions with proteins depends on the type of amino acid residues, their positions relative to each other, and their accessibility in the protein molecule. Deprotonated peptide nitrogen may also coordinate Ni2⫹. In concordance with the highest relative affinity for Ni2⫹ of the histidine imidazole nitrogen and cysteine sulfhydryl group, Ni2⫹ is bound most strongly by the histidyl and also cysteinyl residues in peptides and proteins, and especially by the Xaa-Zaa-His- (or XZH; X and Z stand for any amino acid) motif at the N-terminus. The affinity of Ni2⫹ for His is widely employed for purification of recombinant proteins with N-terminal hexa-histidyl tags, on agarose having immobilized Ni2⫹ ions. Remarkably, the thiol-rich protein metallothionein appears not to be the major Ni2⫹-binding ligand [147]. 2.3.1.1. Nickel Carriers. Transport of nickel in blood plasma is mediated by binding to albumin and some smaller ligands [57,148,149]. The primary nickelbinding site of serum albumin has been identified as the N-terminal XZH motif: e.g., DAHK- in human, DTHK- in bovine, or EAHK- in rat albumin [150,151]. A secondary nickel-binding site involving His-105, His-146, and/or His-247 in the folded molecule, has also been identified in human, bovine, and porcine albumins [152]. The small ultrafiltrable nickel-binding ligands in blood plasma include amino acids and oligopeptides [149,151]. A fraction of plasma nickel is present in nickeloplasmin, which is a nickel-containing α2-macroglobulin [153]. The nickel content of nickeloplasmin is not readily exchangeable with free Ni2⫹, and nickeloplasmin seems not to be involved in the extracellular transport of nickel [154]. In the cytosol of rodent kidney, lung, and liver, parenterally administered nickel was bound to several macromolecular and low-molecular-mass constituents, including GSH and ATP [28, 155–158]. The results from nickel equilibrium studies indicate that nickel–L-histidine complex is the major nickel transporter across the cell membrane, and the nickel–albumin complex serves as the carrier in systemic transport [159]. 2.3.1.2. Regulatory and Structural Proteins. Kondo et al. [160] have found that DAN protein has a nickel-binding motif, PHSHAHPHP, in the C-terminal region. Since DAN has tumor-suppressive activity [161], its possible interaction with cellular nickel may impair cell cycle regulation and assist in carcinogenesis. Met. Ions Life Sci. 2, 621–662 (2007)

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Similarly, cullin-2, a component of the E3 ligase complex that ubiquitinates HIF1α, specifically binds nickel and cobalt at three or more sites [162]. The binding prevents ubiquitination and may thus assist in triggering the hypoxic gene response, typical for these metals. Lipovitellin 2β and importin α appear to be nickel-binding proteins [163, 164] whose interaction with ingested nickel also may deregulate gene expression. A similar binding to neromedin C may impair copper transport and function in the brain [165]. Using nickel-agarose beads, Heiss et al. [166] isolated 22 proteins avidly binding nickel in human B lymphocytes, including tubulin, actin, cullin-2, and at least nine proteins belonging to the heat shock family. Another strong nickel binder is pNiXa protein, the serine protease inhibitor present in Xenopus oocytes and embryos [167]. Disturbance of its function by nickel may result in the observed embryotoxicity and malformation of tadpoles [168]. Xenopus laevis embryos also contain a 40 kDa protein [169], similar to eukaryotic aldolases, that can be isolated on nickel–agarose, and is 96% identical with human aldolase A. Aldolase A may thus also possibly be a target for nickel attack. Nickel and some other transition metals can also bind to the iron regulatory protein-1 (IRP-1), a central regulator of iron homeostasis. Replacement of one iron ion in the 4Fe-4S cluster of IRP-1 with nickel inhibits its enzymatic activity [170] and may contribute to the nickel-induced hypoxic response (see also Chapter 16). Generally, the ability of Ni2⫹ cation to form complexes with a number of proteins raises the possibility that nickel may significantly alter protein conformation and interactions with other proteins, and thus change their functions and cellular homeostasis, producing a variety of pathogenic effects. 2.3.1.3. Histones and Protamines. Binding of Ni2⫹ to DNA is relatively weak, especially in the presence of the physiological DNA counter-ion, Mg2⫹, and amino acid ligands [171]. Therefore, in nuclear and sperm chromatin, the major targets for nickel attack appear to be the proteins, especially the histones and protamines. This subject has been reviewed by Bal et al. [172] and is discussed in Chapter 3. Briefly, strong (at physiological pH) nickel-binding motifs are present in protamine P2 and in core histones H3 and H2A, and weaker ones in histones H2B and H4. Protamine P2 contains the classic XZH N-terminal motif, RTHGQSHYRR-. The histone H3 motif, -CAIH- is located in a hollow structure of the core histone octamer, while the -TESHHKAKGK motif of histone H2A is positioned near the end of its unstructured C-terminal tail. Nickel coordination is also offered by the -AKRHRK- motif in histone H4, [173] and the -ELAKHAmotif in histone H2B [174]. The sequestration of Ni2⫹ by histone tetramer (H3/ H4)2 and histone H2A has been evaluated using numerical models and found to be substantial, even in the presence of maximal physiological concentrations of the major competing cellular ligands histidine and glutathione [158,172,175]. Importantly, Ni(II) coordinated in -CAIH-, RTHGQSHYR-, and -SHHKAKGK Met. Ions Life Sci. 2, 619–660 (2007)


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complexes was capable of mediating oxidative damage to other molecules (see Section 2.3.2). The latter complex is derived from the original Ni(II)-TESHHKAKGK complex of histone H2A owing to a novel effect: nickel-facilitated hydrolysis of the E-S bond [176, 177]. Since the C-terminal tail of H2A is involved in maintaining chromatin structure [178], its hydrolytic truncation in nickelexposed cells may affect chromatin by disturbing orderly gene expression.

2.3.2.

Mediation of Oxidative Damage

The involvement of reactive oxygen species (ROS) in nickel toxicity and carcinogenesis was reviewed previously [179,180]. Soluble and insoluble nickel compounds both produce ROS in cells [181,182]. The oxidative effects of nickel depend on its ability to form the Ni(III)/Ni(II) redox couple around pH 7.4. This is possible only when Ni(II) is complexed by certain natural ligands, including peptides and proteins, especially these which form square planar nickel complexes, e.g., GGH, or GGGG [172,183]. A list of such ligands is provided in [184]. Reactions of such nickel complexes with endogenous O2 or H2O2, generate not only the hydroxy radical •OH (or an oxo-cation NiO2⫹), but also other oxygen-, carbon-, and sulfur-centered radicals originating from the ligands [184–186]. Reactive intermediates can also be produced in the process of oxidative cellular solubilization of nickel sulfides Ni3S2 and NiS. Both are sensitive to oxidation by ambient oxygen, which facilitates their dissolution in biological fluids. The oxidation of Ni3S2 particles is stepwise and eventually leads to intracellular formation of nickel complexes with natural ligands (L), e.g., amino acids and proteins [86]. The first step requires less oxygen: Ni3S2 ⫹1 2 O2 ⫹ 2 HL → 2 NiS ⫹ Ni(II)L 2 ⫹ H 2 O

(1)

Oxidation at this step, initially fast, is slowed by the formation of a protective layer of NiS on the surface of Ni3S2 particles. Further release of nickel requires oxidation of the sulfur moiety of NiS with much more oxygen: NiS⫹ 2 O2 ⫹ 2 HL ⫹ 2 OH⫺ → Ni(II)L 2 ⫹ SO24⫺ ⫹ 2 H 2 O

(2)

In biological fluids, the above reactions are more complex and generate reactive intermediates, e.g., O2 was found to be reduced to H2O2 [187], and sulfur oxidation went through reactive sulfur species [188]. This, obviously, makes nickel sulfides able to generate a wider spectrum of oxidative damage than that produced by other nickel compounds. 2.3.2.1. Lipid Peroxidation. The formation of lipid peroxides results from ROS’s attack on polyenic fatty acid residues of phospholipids (e.g., in cell membranes), Met. Ions Life Sci. 2, 621–662 (2007)

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leading to cell damage. Lipid peroxides decompose to epoxy-fatty acids, alkanes, alkenes, alkanals, alkenals, hydroxyalkenals, and aldehydes, many of which are able to attack and further damage other molecules (reviewed in [189,190]). Nickel and some other transition metals were found to promote lipid peroxidation (LPO) in both in vitro and in vivo experiments. Increased LPO was observed in the brain [121], liver, kidney, lung [106,191,192], and thymus [131] of animals given parenteral nickel; and in isolated lymphocytes [193], blood platelets [194], rat kidney epithelial cells [189], and liver microsomes [195], treated with nickel in vitro.

2.3.2.2. Protein Damage. The oxidation of amino acids and proteins by ROS and the roles of toxic metals in this process have been reviewed before [196,197]. Protein oxidation is mechanistically involved in many adverse effects, including cancer [184–186,196,197]. Nickel, like many other transition metals, may promote oxidative modification of both free amino acids and the amino acid residues in proteins. In proteins, major targets are the side chains of Cys, His, Arg, Lys, and Pro. The sulfhydryls are commonly oxidized to disulfides, but may also be converted into sulfino-, sulfeno-, and solfono- derivatives; this is also true for the Met residue. His imidazole may be oxidized to aspartic acid, asparagine, or 2′-OH-His. Arg converts into γ -glutamic-semialdehyde; Lys, to 2-amino-adipicsemialdehyde; and Pro into glutamic acid, pyroglutamic acid, γ -aminobutyric acid, and γ -glutamic-semialdehyde [1,4,10,198]. Glutamic and aminoadipic semialdehydes have been identified as the main carbonyl products of metal-catalyzed oxidation of proteins [197]. The formation and rearrangements of radical intermediates arising in the oxidation process of proteins may lead to protein fragmentation and intra- and inter-protein cross-linking. The proteins may also cross-link with DNA [184– 186]. Because of the site-specific nature of nickel binding, ROS generation and protein damage are also site-specific. Therefore, nonbinding residues, such as Trp, Tyr, Phe, and Met, also sensitive to ROS attack, may not be targeted by the metal-catalyzed oxidation [196]. They can, nonetheless, be damaged if they are located close to the metal binding site [199]. A good example of both oxidative and conformational effects of Ni(II) on polypeptides is interaction of the latter with a 15-mer peptide, RTHGQSHYRRRHCSR-amide (HP21-15), modeling the N-terminal sequence of human protamine P2 [199–201]. When bound to the RTH- end motif of this peptide, nickel catalyzes oxidation by H2O2 of not only Arg-1 and His-3, but also Tyr-8. The reason for this is a strong structuring effect of nickel on the peptide ligand that brings Tyr-8 close to the metal center [199] and shifts all the positive Arg side chains to one side of the molecule. Thus, by imposing conformational changes on its ligand, the bound Ni(II) directs oxidative damage to particular amino acid residues. It also modulates the ligand function by increasing its binding with DNA [199–201]. Met. Ions Life Sci. 2, 619–660 (2007)


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The coordination mode was the most likely cause of a profound difference in redox activity between nickel complexes originating from the C-terminal ‘tail’ of histone H2A. Nickel is bound to the -TESHHK- motif of this tail to form a non-redox-active octahedral complex [176,202]. However, the binding is followed by hydrolysis of the E-S peptide bond with formation of the SHHKAKGK peptide. The latter binds nickel very strongly through the SHH- motif, yielding a square planar complex [203] that is redox-active at physiological pH. Its reaction with H2O2 results in oxidation of the Ser and His residues of -TESHHK- and collateral oxidative damage to DNA, if added to the reaction mixture [176].

2.3.2.3. DNA Damage. In chromatin, metal cations are offered an abundance of binding sites by both DNA and proteins. In effect, following in vivo exposure, nickel and other heavy metals are found in cell nuclei [98,204–206]. Since, however, DNA is blocked by the physiological cation Mg2⫹, the toxic metals are likely to associate predominantly with histones, the prevalent nuclear proteins [184,186,200,202,207]. Unlike nickel bound to DNA [208,209], the protein-bound nickel can catalyze ROS generation and thus induce oxidative damage to all chromatin components, including DNA. Generally, oxidative effects in DNA observed in animals and cultured cells exposed to nickel are cross-links of various types, strand scission, depurination, and base modifications [10,98,184–186,206,210]. The most common effect of nickel and other toxic metals in chromatin observed in vitro and in vivo is DNA–protein cross-linking [10,98,184–186,206,210,211]. Metal ions can induce DNA–protein cross-links in two ways: through the formation of mixed-ligand complexes, or by assisting in generation of strong covalent bonds directly between DNA and the proteins. In the case of nickel, the formation of cross-links of both types has been observed in vitro [188,212] and in vivo [213]. Intra- and inter-strand cross-linking in DNA is also possible, but the effect of nickel on its formation has been observed only indirectly, thanks to tandem double CC → TT mutations typical for cross-links between cytosines in the same strand. Mutations of this type arose on template DNA pre-treated with nickel and hydrogen peroxide [214]. The presence of cross-links in chromatin may lead to morphologic aberrations of chromosomes, as observed in lymphocytes of workers exposed to nickel compounds [184–186]. In cultured CHO cells, chromosomal aberrations caused by nickel were predominantly localized in their heterochromatic regions [215]. Another common ROS effect is DNA cleavage. Thus, single-strand breaks were found in kidneys and lungs, but not in livers of rats after parenteral administration of nickel chloride [216]. In vitro, single- and double-strand breaks were observed in soluble nickel-treated blood lymphocytes [217] and in HeLa cells cultured with Ni3S2, but not with NiO [218]. Single-strand breaks also were produced in isolated DNA by soluble nickel plus H2O2 [184–186,210,211]. This effect was enhanced by peptides forming square planar nickel complexes [200,201,219]. Met. Ions Life Sci. 2, 621–662 (2007)

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Kawanishi et al. [220] have found that nickel promotes in vitro DNA cleavage by H2O2 in a site-specific way. The most sensitive sites appeared to be at the cytosine, thymine, and guanine residues. DNA cleavage mediation by nickel and other metal complexes, and ligand effects on the selectivity of DNA oxidation with various oxidants were studied in detail by several laboratories (reviewed in [184–186]). The spectrum of damage produced by ROS in the base moiety of DNA is wellestablished [221]. Many modified DNA bases have been found in isolated chromatin or DNA exposed to H2O2 plus nickel, cobalt, copper, and iron salts. The most abundant is usually 8-oxoguanine. Under ambient O2, nickel assisted in generation of such bases in chromatin, but not in pure DNA, indicating facilitation of its redox activity by chromatin proteins, most likely the core histones [209]. Besides attacking the bases directly in DNA, ROS may damage the nucleobases in triphosphonucleosides from which the bases may be misincorporated into genomic DNA, or RNA, and cause mutations [222]. Following in vivo exposure to soluble nickel, elevated amounts of at least one damaged DNA base were found in organs of rats [213,218,223,224] and mice [184]. In the lungs of rats, both insoluble Ni3S2 and NiO and soluble nickel sulfate, instilled intratracheally, increased pulmonary 8-oxo-dG levels, but in cultured HeLa cells, only Ni3S2 was active. This difference seems to indicate different mechanisms of the damage in vitro and in vivo by the same compounds, perhaps owing to contribution of ROS generated by inflammatory cells in the rat lung [218]. In mice treated with soluble nickel, renal 8-oxo-dG levels were increased only in the BALB/c strain, which had low glutathione and glutathione-peroxidase levels compared with two other strains, B6C3F1 and C3H [184,225]. DNA damage resulting from exposure to toxic metals also includes depurination. Nickel produced apurinic sites in DNA and mediated in vitro hydrolysis of 2′-deoxyguanosine [226,227]. The depurination occurred concurrently with DNA strand scission; both effects could result from ROS attack on DNA sugar moiety. Modified sugars constitute alkali-labile sites found in DNA from metaltreated cells [184–186,226,227].

2.3.2.4. Effect on Ascorbic Acid, Glutathione, and Other Antioxidants. The oxidative damage, induced by nickel, may be aggravated by a concurrent depletion by nickel of major cellular antioxidants, such as ascorbate and glutathione, or inhibition of antioxidant enzymes. The affinity of ascorbic acid for nickel is low [228]. Nonetheless, nickel can deplete intracellular ascorbate [229] by catalytic oxidation [230,231] and inhibition of the sodium-dependent ascorbic acid transporters SVCT1 and SVCT2 [232] (own unpublished results). The depletion leads to the inhibition of cellular hydroxylases that, in turn, may result in activation of the HIF-1-dependent hypoxic response, impairment of assembly of proteins with collagen-like domains, and/or inactivation of certain ascorbate-dependent DNA repair enzymes [229,231]. Ascorbic acid supplementation averts the hypoxic Met. Ions Life Sci. 2, 619–660 (2007)


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response to nickel [229]. Ascorbic acid alleviated the toxicity of nickel and other heavy metals in animals, cultured cells [194,233–236], and humans [237]. Thus, vitamin C supplements might prevent metal toxicity. However, in developing preventative measures, we must also consider data from some in vitro experiments in which ascorbic acid aggravated metal toxicity [226,238], consistently with the observed generation of ROS in the process of metal-catalyzed ascorbate oxidation [239]. In a test-tube, glutathione, another natural antioxidant, can form complexes with transition metals in both reduced (GSH) and oxidized (GSSG) forms. Krezel et al. recently described the Ni(II)-GSH complex system [158,240], noting acceleration of Ni(II)-GSH oxidation in alkaline, but not in neutral solutions. However, as indicated by the stability constants of respective complexes, in the cellular environment, histidine and ATP rather than GSH, may prevail as ligands for nickel. Thus, a direct impact of nickel on GSH level and its antioxidant function may be limited. It concurs with other observations of limited inhibition by GSH of nickel toxicity, especially that mediated through ROS generation [241]. The depletion of GSH observed in cells exposed to nickel [190,242] would thus be a secondary effect due to scavenging by GSH of nickel-generated ROS. The importance of GSH in the defense against ROS has also been indicated by a marked increase of GSH production in cells resistant to nickel and hydrogen peroxide [182], and activation of the GSH/GSSG related enzymatic system in cells exposed to low doses of nickel [243]. Interestingly, the cytoplasmic and mitochondrial thioredoxins, also considered as part of the thiol antioxidant system, appeared to be less responsive to nickel than GSH [242]. The enzymatic components of cellular antioxidant defense may also be affected by nickel. In vitro, nickel at millimolar concentrations inhibited enzymatic activity of superoxide dismutase (SOD) [244] and catalase (CAT) [245]. In vivo, nickel effects on SOD, CAT, glutathione peroxidase (GSH-Px), and glutathione reductase (GSSG-R) appear to be indirect. They include either activation or inhibition of a particular enzyme, depending on the animal and target cell, nickel dose, and the regimen of exposure. For example, the same dose of nickel chloride lowered hepatic GSH level in mice, but not in rats; while in guinea pigs, a significant increase was observed [246]. A similarly uneven pattern of changes among SOD, CAT, and GSH-related enzymes (GSH-Px, GSSG-R, glutathione S-transferas, and γ -glutamyl transferase) was observed in the liver, kidney, and skeletal muscle of rats given a high (107 µmol/kg) intraperitoneal dose of nickel acetate [247]. However, a much lower, sub-micromolar intratracheal dose of nickel chloride increased SOD activity in the rat lung for almost 40 days [248]. The inhibition by nickel of the major enzymatic antioxidants, CAT, GSH-Px, GSSG-R, or SOD, observed under certain conditions, may enhance nickel-induced oxidative damage. A similar enhancement may result from stimulation by nickel of myeloperoxidase release from polymorphonuclear lymphocytes in nickel dermatitis [249]. Met. Ions Life Sci. 2, 621–662 (2007)

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3.


NICKEL-INDUCED CARCINOGENESIS

3.1.

Human Epidemiology

The development of cancers of the nasal cavities in Welsh nickel refinery workers was first reported by Bridge in 1933. In 1937, Baader described 17 nasal and 19 lung tumors among workers of the same refinery. By 1949, these numbers increased to 47 nasal and 82 lung cancers, and cancers at both locations were proclaimed in Great Britain as industrial diseases among nickel workers [5,250]. Following these pioneering findings, the carcinogenicity of nickel compounds has been confirmed by numerous epidemiological studies in humans and experimental bioassays in animals [3–5,8,251–253].

3.1.1.

Carcinogenic Nickel Species

Epidemiology pointed at increased mortality from respiratory tract malignancies in refinery workers chronically exposed to nickel-containing dusts and fumes. Welding of nickel alloys also may be a source of such fumes [8]. For many years, it was believed that only water-insoluble nickel components of the dusts (e.g., Ni3S2, NiO) were carcinogenic. However, more recent data clearly indicate that aerosols of water-soluble nickel compounds generated in nickel electro-refining plants are carcinogenic to humans as well, with a clear dose-related effect [3,251]. Tobacco smoking was found to be only a weak to moderate confounder [3]. In addition to lung and nose tumors, increased risks of occupation-related malignant tumors of the larynx, kidney, prostate, stomach, and soft-tissue sarcomas, have been reported, but the significance of these findings is doubtful. Also, no clear evidence exists of possible cancer risk from general environmental and dietary nickel [254]. Besides occupational exposures, nickel present in endoprostheses, bone-fixing plates and screws, and other medical devices made of metal alloys, has been suspected, but not proven, to be the cause of sporadic local tumors [14,255,256]. The carcinogenic effects of metallic nickel and nickel compounds have been evaluated by the International Agency for Research on Cancer (IARC), which concluded that nickel sulfate and the combinations of nickel sulfides and oxides encountered in the nickel refining industry are carcinogenic to humans (Group 1) and that metallic nickel is possibly carcinogenic to humans (Group 2B) [8].

3.1.2.

Histopathology of Nickel Cancer

Histopathology of the human respiratory tract tumors was compiled by Sunderman et al. [257]. Among the sinonasal cancers were squamous cell carcinomas (48%), anaplastic and undifferentiated carcinomas (39%), adenocarcinomas (6%), Met. Ions Life Sci. 2, 619–660 (2007)


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transitional cell carcinomas (3%), and other malignant tumors (4%). Lung tumors were diagnosed as squamous cell carcinomas (67%), anaplastic, small cell, and oat cell carcinomas (15%), adenocarcinomas (8%), large cell carcinomas (3%), other malignant tumors (1%), and unspecified cancers (6%). Thus, prevalence of squamous cell carcinomas was evident.

3.2. 3.2.1.

Carcinogenesis in Experimental Animals Carcinogenic Nickel Compounds

Campbell was the first to report in 1943 that inhalation of nickel dust caused a two-fold increase of lung tumor incidence in mice [258]. Since then, experiments in animals have revealed carcinogenicity of nickel compounds with low or no aqueous solubility (e.g., Ni(OH)2, Ni3S2, NiO) following inhalation or parenteral administration. Carcinogenesis of soluble nickel compounds was studied less extensively, but also yielded positive results after parenteral [259–262] or transplacental injections [263]. As a rule, however, insoluble compounds are thought to be stronger carcinogens than soluble compounds, most likely because of longer retention at the target tissue [264,265].

3.2.2.

Administration Routes

Nickel compounds induce tumors at virtually all application sites (reviewed in [8,10,253]). The administration routes tested in animals include inhalation and intramuscular, intrarenal, intraperitoneal, intraocular, subcutaneous, and intraarticular injections. 3.2.2.1. Intramuscular and Subcutaneous Injections. The development of highly malignant rhabdomyosarcomas in mice and rats following intramuscular injection of water-insoluble Ni3S2 or NiO was first reported by Gilman in 1962 [266]. Since then, his experimental model has been used in numerous experiments in various animal species and strains, as reviewed in [267]. Experiments with 63Ni and 35S labels showed that the carcinogenic Ni3S2 particles persisted at the injection site for months [85]. A high incidence of local rhabdomyosarcomas and fibrosarcomas was also observed in rats after subcutaneous injection of Ni3S2 [268,269]. 3.2.2.2. Intraperitoneal Injection. Increased lung tumor incidence was noted in strain A mice after multiple intraperitoneal injections of nickel acetate[270]. Low incidence of renal cortical adenomas was observed in male F344 rats after a Met. Ions Life Sci. 2, 621–662 (2007)

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single intraperitoneal injection of nickel acetate, followed by a prolonged dietary treatment with tumor promoter sodium barbital [261]. Intraperitoneal injection of nickel acetate to pregnant F344 rats resulted in tumors in the offspring: pituitary gland tumors developed without barbital, and renal tumors occurred only when nickel was followed by barbital administration. These results indicate that nickel acetate is a complete carcinogen for fetal rat pituitary gland and a potent initiator of carcinogenesis in fetal rat kidney [263]. 3.2.2.3. Intrarenal Injection. Kidney tumors developed following intrarenal injection of Ni3S2 in different strains of rats [271–273]. No such cancers were observed in rats injected with metallic nickel or NiS dust. Attempts to induce tumors by intrarenal injection in other animals were not successful [272]. Histologically, most of the rat renal tumors resembled the sarcomatous variant of the classic renal mesenchymal tumor, while some were composed of poorly differentiated cells [114,273]. Interestingly, the early stage of renal carcinogenesis by Ni3S2 was accompanied by a strong increase in hemoglobin and erythrocyte levels [114,274]. The erythrocytosis was due to the induction of erythropoietin [111,275], a part of the hypoxia-mimicking response to nickel, reviewed in more detail in Chapter 16. 3.2.2.4. Intratesticular Injection. Malignant testicular tumors developed in 16 of 19 rats within 20 months after a local injection of Ni3S2. They were classified as fibrosarcomas, malignant fibrous histiocytomas, and rhabdomyosarcomas. Since rhabdomyosarcomas are not typical for the testis, the authors have suggested that Ni3S2 induces malignant transformation of undifferentiated, pluripotent mesenchymal cells [276]. 3.2.2.5. Intraocular Injection. Intraocular injections of Ni3S2 to rats resulted in the development of local malignant tumors [277]. It is of interest that Ni3S2 can also induce ocular tumors in an evolutionary distant species like the Japanese newt, Cynops pyrrhogaster [278]. 3.2.2.6. Inhalation. Oller et al. [252] published a review of inhalation carcinogenesis by selected nickel compounds. Early on, lung tumors were induced in rats following inhalation of Ni(CO) 4 vapor [279], Ni3S2 dust [280], or nickel feinstein dust [281]. Clear evidence for lung tumorigenicity of Ni3S2 dust in rats after inhalation for more than one year was presented in another study [282]. Likewise carcinogenic was insoluble nickel oxide in a large inhalation bioassay conducted by the National Toxicology Program in rats [101]; in contrast, however, soluble nickel sulfate was found not to be carcinogenic [99]. Mice were more resistant to nickel inhalation carcinogenesis than rats [282]. Met. Ions Life Sci. 2, 619–660 (2007)


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A single intratracheal instillation of Ni3S2 resulted in the development of only one tumor in 26 exposed rats, likely because of fast clearance of the deposited particles [281]. However, Ni3S2 readily induced carcinomas and sarcomas in the epithelium of heterotopic tracheal transplants, which could not eliminate the carcinogen [283]. 3.2.2.7. Other Routes of Exposure. There is no evidence of cancer induction by soluble nickel administered in drinking water. Also, no tumors were observed following treatment of hamsters with insoluble Ni3S2 by multiple applications into the cheek pouch, oral cavity, or the digestive tract, in total dosages as high as 1 g of Ni3S2. Likewise, no malignant tumors developed in rats that received single injections into the submaxillary gland or into the liver [284].

3.2.3.

Species and Strain Susceptibility

Although no absolute species specificity has been observed in nickel carcinogenesis, rats are apparently more susceptible than mice, hamsters, or rabbits [285]. Also, significant variations in susceptibilities have been noticed within rat and mouse strains [285,286]. For example, the incidence of renal tumors after intrarenal injection of Ni3S2 in different strains of rats ranged from 64% in Wistar– Lewis rats, 50% in NIH Black rats, 28% of F344 rats, to none in Long–Evans rats [271–273]. Most rat organs, including brain [287], are susceptible to nickel carcinogenesis via local administration. Intraocular and intramuscular injections yield the highest tumor incidences. The strain differences in rats might depend on different abilities of phagocytes to ingest nickel particles [288]. It is also possible that genetic differences between animals resulting in various levels of activity of antioxidant systems influence the response to nickel [106,286]. Differences in carcinogenic activity of nickel compounds between rats and mice were also evident after inhalation or tracheal instillation. Intratracheal Ni3S2 failed to induce tumors in B6C3F1 mice [289], while the same compound induced adenomas and carcinomas in 30% of exposed F344 rats [102]. However, nickel-caused inflammation and lung fibrosis developed in both species.

3.2.4.

Interactions with Other Carcinogens

Administration of nickel compounds with organic carcinogens produced a significant synergistic effect. A single intramuscular injection of Ni3S2 with 3,4benzopyrene to rats induced more sarcomas and in a much shorter time than injection of the carcinogens alone [290]. A similar effect was observed after intratracheal instillation of the same compounds to rats [291]. Most of the muscle sarcomas caused by nickel alone or in combination with 3,4-benzopyrene were classified as rhabdomyosarcomas that are typical for nickel in rats, whereas exposure to 3,4-benzopyrene alone produced fibrosarcomas [290]. Met. Ions Life Sci. 2, 621–662 (2007)

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3.2.5.


Interactions with Essential Metals

The essential metals Mn(II) [292], Mg(II) [132,293,294]], and Zn(II) [294], but not Ca(II) [293], co-administered intramuscularly with Ni3S2 to rats, reduced local tumor incidence in a dose-dependent manner. Magnesium was the strongest and zinc the weakest inhibitor. However, separate parenteral or dietary administration of the essential metals did not produce this effect. Magnesium carbonate co-administered with Ni3S2 was quickly absorbed from the muscle (hours) and had no effect on the retention of Ni3S2 at the injection site (weeks) [293]. Thus, magnesium inhibited nickel action only at the initiation stage of carcinogenesis, thus revealing the multi-stage character of Ni3S2 carcinogenesis. Magnesium also strongly inhibited renal carcinogenesis by Ni3S2 in the rat [114]. Co-injection of Ni3S2 with iron, as either metallic powder (Fe0) or Fe(III) sulfate, strongly inhibited Ni3S2 carcinogenicity in the rat muscle [295]. In contrast, when co-injected with Ni3S2 into the rat kidney, Fe0 did not affect the final yield of tumors, but significantly shortened their latency. Neither Fe0 nor Fe(III) induced muscle or kidney tumors by themselves [114]. In strain A mice, multiple intraperitoneal injections of nickel acetate with magnesium or calcium acetates resulted in a lower incidence of lung adenomas than that produced by nickel alone [260].

3.3. Malignant Transformation of Cultured Cells Soluble and insoluble nickel compounds are capable of transforming cultured human and rodent fibroblastic and epithelial cells [296]. In rodent cells, which transform more easily than human cells, the insoluble compounds act like complete carcinogens. For example, in Syrian hamster embryo (SHE) cells, Ni3S2 induced morphological transformation, growth in soft agar, and development of tumors upon injection to nude mice [297]. In the same cells, soluble nickel was less potent and produced only fast-growing immortalized colonies [298], although at equitoxic doses, insoluble Ni3S2, NiO, Ni2O3, and soluble nickel acetate showed equal transforming effects in BHK-21 cells [299]. The ability of soluble nickel to immortalize cultured cells was found to be higher than that of other carcinogens, including benzo[a]pyrene diol epoxide, N-methyl-N-nitrosourea or, γ or X-rays [298]. The exposure of mouse C3H/10T1/2 cells to Ni3S2 caused morphological transformations [300]. Primary human kidney epithelial cells cultured with soluble nickel became immortal, grew in soft agar, and developed abnormal karyotypes, but did not grow in nude mice [301]. Tumorigenicity was achieved in these cells only through transfection of activated Ha-ras oncogene [302]. In addition to the cell-transforming potential, typical for tumor initiators and complete carcinogens, nickel also has the properties of tumor promoters. These were observed in SHE [303] and in NIH 3T3 cells [304]. Met. Ions Life Sci. 2, 619–660 (2007)


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Mechanisms of Nickel-Induced Carcinogenesis Genotoxic Effects

Nickel compounds induce chromosomal damage [305]. This has been especially notable in the heterochromatic long arm of the Chinese hamster X chromosome, which showed regional decondensation, frequent deletions, and other aberrations following treatment of CHO cells with nickel [306]. Also in CHO cells, nickel compounds induced sister chromatid exchanges (SCE), especially in heterochromatin [307]. In cultured human lymphocytes, nickel sulfate induced a nearly twofold increase in SCEs [308], while Ni3S2 increased the frequency of micronuclei formation [309]. Other symptoms of nickel genotoxicity included microsatellite mutations in human lung cancer cells [310], insertion mutations in rat kidney epithelial (NRK) cells infected with MuSVts110 retrovirus [311], and deletion mutations in CHO cells [312]. The G → T transversion, typical for oxidative DNA damage, was found in the K-ras gene (codon 12) in renal tumors induced by Ni3S2 alone or combined with iron powder [313]. The same type of point mutation in the p53 gene was associated with nickel-exposure-related human lung tumors [314]. Despite numerous reports of the DNA and chromatin damage in nickelexposed mammalian cells and tissues, the results of mutagenic assays in bacteria, fruit fly, and some mammalian cells [96, 315–320] indicate a low mutagenic potential for this metal. Nickel was not mutagenic in the S. typhimurium and E. coli test systems [315, 316], but did mutate Corynebacterium sp. 887 (hom) [316], and could act as a potent co-mutagen with alkylating agents in some E. coli and S. typhimurium tester strains [321]. So, the effect depends on the model used. More on this subject may be found in Chapter 16.

3.4.2.

Epigenetic Effects

Epigenetic toxicity of nickel that may lead to cell transformation is mainly realized through deregulation of gene expression, as discussed in detail in Chapter 16. Molecular events that may lead to such deregulation are reviewed below.

3.4.3.

Molecular Basis for the Genetic and Epigenetic Toxicity of Nickel

3.4.3.1. Promutagenic Effects of Oxidative DNA Damage. The strongest association of oxidative damage with carcinogenesis stems from the mispairing and thus promutagenic properties of many DNA base products generated by ROS and found in nickel-exposed cells in vitro and in vivo. Other DNA lesions, such as strand scissions and depurination, which have also been found, may lead to mutations, too [184–186,226,227]. Therefore, the formation of these lesions in Met. Ions Life Sci. 2, 621–662 (2007)

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DNA may be considered as a genotoxic event leading to cell death or mutation. Most importantly, nickel can also inhibit DNA repair systems [322,323]. However, mutations are not the only effect produced by the damaged DNA bases. 8-Oxoguanine, for example, may also misdirect DNA methylation [324] and perturb orderly binding of transcription factors to DNA [325]. These effects may assist tumor promotion and progression. Physiologically generated ROS serve as signal transduction messengers in controlling gene expression [186,326]. Therefore, it seems likely that the increase in redox reactions driven by intracellular nickel may disturb orderly generation of messenger ROS and affect redox-regulated proteins, such as NF-κ B, AP-1, p53, K-ras, Bcl-2, HIF-1, and others (see Chapter 16). This must disturb progression of the cell cycle and/or apoptosis. The reduced level of binding of NF-κ B and AP-1 transcription factors to DNA in nickel-resistant transformed cells, and their vigorous response to H2O2 or buthionine sulfoximine, clearly indicate that nickel resistance is closely allied to oxidative stress responses [182]. Nickel-mediated attack on regulatory proteins not belonging to the redox signaling network may affect their structure and function too, as exemplified by the Fhit protein inhibition [327]. Also, binding of nickel to histones may sensitize them to oxidation and other types of damage (see Sections 2.3.2.2. and 3.4.3.3.) and eventually lead to changes in chromosome morphology and gene expression [328]. 3.4.3.2. Inhibition of DNA Repair Enzymes. Inhibition of any element of the DNA repair systems may potentially assist in cell mutation and carcinogenesis. In vitro, nickel was found to impair the function of DNA polymerase and cause base misincorporation into newly synthesized oligonucleotides [329]. Extensive investigations by Hartwig et al. revealed that nickel, like some other carcinogenic metals, did inhibit base and nucleotide excision repair. Such inhibition enhanced DNA damage by UV radiation, ROS, benzo[a]pyrene-7,8-diol 9,10-epoxide, and methylating agents [322,330–333]. The proteins targeted by nickel in DNA repair machinery include the XPA zinc-finger protein [332,334] and O6-methylguanineDNA methyltransferase [333], but not Fpg-glycosylase [335]. Effects of nickel and other metals were also tested on the bacterial and human 8-oxo-dGTPases, MutT and MTH1. These enzymes prevent potentially mutagenic utilization of 8-oxo-dGTP and other damaged nucleoside triphosphates for DNA synthesis (reviewed in [336]). 3.4.3.3. Effects on DNA Methylation and Histone Modifications. Nickel enhances DNA methylation and affects acetylation, methylation, and other modifications of histones in cultured cells. These effects are targeted toward DNA and histones at certain chromosomal regions and specific histone sites. They may modify chromatin structure and gene expression, and thus assist in cell transformation through epigenetic mechanisms (see Chapter 16 for more details). Met. Ions Life Sci. 2, 619–660 (2007)


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Downregulation by nickel of histone acetylation has been reported for both yeast and mammalian cells [337,338]. This effect was most likely mediated through inhibition of specific histone acetyltransferases [337]. In human and rat cell lines exposed to nickel, acetylation of all four core histones was reduced in a dose- and time-dependent way. Acetylation of histone H2B was suppressed to the highest and that of histone H3 to the lowest extent. In H2B, the decrease of acetylation was site-specific [338]; the same was observed in yeast histone H4 [337]. A decrease in histone acetylation is expected to inhibit gene expression. However, in cells exposed to nickel this was not always the case [338–340], indicating a complex dependence, probably due to the site-specificity of nickel effect. Analysis of core histones extracted from nuclei of cells exposed to nickel also revealed changes in their ubiquitination levels [341] and the presence of certain unique structural modifications, such as truncation and deamidation [342,343]. The truncation afflicted histones H2A and H2B. In the case of H2A, the cut-off peptide, SHHKAKGK, was identical with that observed in test-tube experiments [176, 202], implicating a direct nickel-assisted hydrolysis at the C-terminal tail of this histone. In contrast, histone H2B was truncated on both termini in two identical -KAVTK- repeats by specific proteolytic enzymes, calpains. Besides being truncated, histone H2B from nickel-treated cells also had its Gln-22 residue deamidated and Met-59 and Met-62 residues oxidized to sulfoxides, a signature of oxidative stress [343]. Since the enzymatic activity of calpains depends on calcium, the truncation of H2B is yet another symptom of the disturbance by nickel of calcium homeostasis. 3.4.3.4. Disruption of Calcium Homeostasis. Nickel blocks calcium channels [344] and deregulates intracellular calcium homeostasis [345]. One result of altering intracellular calcium metabolism by nickel was rapid proliferation of nickel-transformed cells in low-calcium media [346]. Since cytoplasmic Ca2⫹ pulses regulate expression of genes associated with growth, differentiation, and apoptosis of cells [347,348], one can presume that derangement of such pulses may have pathogenic consequences, including malignant cell transformation. These data and Jaffe’s ‘calcium theory of oncogenesis’ (reviewed in [171,349]) prompted some broader investigations of nickel/calcium interactions in gene expression signaling discussed in Chapter 16.

4.

CONCLUSION

Nickel is ubiquitously present in the environment and intake of low doses of its compounds is unavoidable. This may be harmless to the general population, but not for individuals allergic to nickel. Occupational exposures, such as those in Met. Ions Life Sci. 2, 621–662 (2007)

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nickel refining, electroplating, and welding, have the potential to provide nickel doses high enough to induce pathologic reactions, including allergic dermatoses and cancer of the respiratory tract. As indicated by experimental data, the toxicity and carcinogenicity of nickel stem from its potential to damage proteins, nucleic acids, and smaller molecules [176,184–186,350–353]. The type and extent of the damage depend on the intracellular dose of nickel ions that, in turn, is a function of physicochemical properties of particular nickel derivatives, their ability to enter the cell and/or to dissolve within the cell. Therefore, lipid-soluble nickel carbonyl has the highest, and insoluble nickel-containing dusts have the lowest acute toxicities. Because of limited cellular uptake, water-soluble nickel compounds are neither highly toxic nor carcinogenic unless administered parenterally. The wide spectrum of molecular toxicity of nickel results from redox activity of its complexes with certain cellular ligands at physiological pH. Such ligands include some amino acids, peptides, proteins, and other molecules, but not DNA [184–186,208,209,354]. ROS generated in reactions of these complexes with dioxygen and its metabolic derivatives (•O⫺2 , H2O2, lipid peroxides) are capable of inflicting both intrinsic and collateral damage to biomolecules, e.g., proteins and DNA. The exceptionally high carcinogenic potential of Ni3S2 among nickel compounds originates from redox reactions involving both nickel and sulfur. In addition, the capacity of nickel to inhibit DNA repair [322,330–333] may contribute to the persistence of oxidative DNA damage caused by ROS. The oxidative damage to proteins and nucleic acids can produce lethal, mutagenic, and epigenetic effects. In humans and animals, nickel attacks the immune system, activating the inflammatory response and increasing oxidative stress [218]. In addition, inhibition of the natural killer cells by nickel [132] may suppress elimination of mutated cells. The strongest epigenetic effects of nickel have been associated with the hypoxic gene response caused by inhibition of HIF-1α hydroxylation (see Chapter 16). HIF-1 upregulates numerous genes involved in glucose transport and glycolysis [355]. Therefore, chronic nickel exposure is likely to promote selection of cells maintaining a high glycolytic rate typical for cancer cells, as first described by Warburg [356]. The mechanisms of HIF-1 activation by nickel involve depletion of cellular ascorbate with the resulting inactivation of prolyl hydroxylases. Proline hydroxylation is crucial not only for regulation of HIF-1, but also for proper assembly of collagen, extracellular matrix, and other proteins having collagen-like domains, such as the lung surfactant. Hence, the depletion of ascorbate can be especially deleterious for lung cells, the major target for nickel toxicity and carcinogenesis [231]. Since the inhibition by nickel of HIF-1α prolyl hydroxylase is reversed by ascorbic acid supplementation [229], vitamin C could possibly be used to prevent nickel toxicity and carcinogenesis in humans. Therefore, there is an urgent need for more research in this field. Also, nickel toxicology greatly benefits from in-depth understanding of nickel interactions with bioligands and essential metals. Research in this field is therefore equally urgent. Met. Ions Life Sci. 2, 619–660 (2007)


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ACKNOWLEDGMENTS This work was supported by the Intramural Research Program of the NIH, NCI.

ABBREVIATIONS CAT CHO cells dithiocarb DTPA GSH GSH-Px GSSG GSSG-R HIF IARC IRP LD50 LPO NK ROS SCE SHE cells SOD

catalase Chinese hamster ovary cells diethyldithiocarbamate diethylenetriaminepentaacetic acid reduced glutathione glutathione peroxidase oxidized glutathione glutathione reductase hypoxia-inducible factor International Agency for Research on Cancer iron regulatory protein lethal dose of a substance, which leads, given in a single dosis, to the death of 50% of a population lipid peroxidation natural killer reactive oxygen species sister chromatid exchange Syrian hamster embryo cells superoxide dismutase

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351. 352. 353. 354. 355. 356.

Met. Ions Life Sci. 2, 619–660 (2007)

Subject Index

A A. cylindrica, see Anabaena eutrophus, see Alcaligenes fulgidus, see Archaeoglobus japonicus, see Aspergillus metalliredigenes, see Alkaliphilus niger, see Aspergillus pleuropneumoniae, see Actinobacillus thaliana, see Arabidopsis vinosum, see [NiFe] hydrogenases AAS, see Atomic absorption spectroscopy Absorption of nickel, 621, 622 Absorption spectroscopy nickel superoxide dismutase, 433 ACDS, see Acetyl-coenzyme A decarbonylase/synthase Acetate (or acetic acid) bridge, 219 buffer, 161 cleavage, 359, 388 2-cyano-2-(hydroxyimino)-, 75, 76 degradation, 385 formation, 359, 386, 388 nickel(II), 604, 627, 628, 637, 639, 640, 642 oxidation, 359 reduction, 384


Acetohydroxamic acid, 228 urease complex, 259–261, 272 Acetonitrile as ligand, 218 Acetonylphosphonate Ni(II) complex, 117 Acetyl-coenzyme A biosynthesis, see Biosynthesis formation, 389, 390, 393 Acetyl-coenzyme A decarbonylase/ synthases, 326, 327, 369, 376, 383–386 archaeal, 386–389 evolution, 386–393 genome sequences, 409, 410 physiological roles, 386–393 properties, 359, 360 subunit phylogeny, 371, 372, 381, 382 Acetyl-coenzyme A synthases, 376, 383, 524, 530 active site, see Active site mechanism, 378 models, 200–212 molecular chaperones, 538 synthesis, see Biosynthesis Acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 35, 37, 81, 182, 360, 366, 369, 371, 373–381, 385, 524, 525 acetylated state, 379, 380 A-cluster, see Clusters


662

[Acetyl-coenzyme A synthases/carbon monoxide dehydrogenases] Carboxythermus hydrogenoformans, 359 catalytic cycle, 379, 380 conformational change, 381, 382 conserved residues, 375 evolution, 392, 393 genes in bacteria, 373, 389–392, 538 inhibition, 381 methylated state, 379, 380 models, 200–211 Moorella thermoacetica, 359, 361, 365 Ni(0) state, 377–379 Ni(I) state, 204, 205, 377–379 properties, 359 protein sequence alignments, 402–404 reductive activation, 377–379 structure, 362 subunit phylogeny, 371–373, 381, 382, 391 Acetyl phosphate Ni(II) complex, 115–117 Acidity constants (see also Equilibrium constants) of 5⬘-ATP, 143 adenosine, 151 amino acids, 67 buffers, 163 dihydroxyacetone phosphate, 117 1,N6 -ethenoadenosine nucleotides, 150 flavin mononucleotide, 148 glycerol 1-phosphate, 117 hydroxamic acid, 72, 73 methane, 349 methyl thiophosphate, 155 microconstants, 73, 120, 137, 138, 155 Ni(aq) 2⫹, 111 nucleoside 5⬘-diphosphates, 142 nucleoside 5⬘-triphosphates, 143 nucleotides, 115, 156 phosphate groups, 128, 129, 148, 158 purine nucleobases, 119, 120, 159 purine-nucleoside 5⬘-monophosphates, 135, 146 purine-nucleotide (N1)-oxides, 152–154

SUBJECT INDEX

[Acidity constants] pyridine, 120 pyrimidine-nucleoside 5⬘monophosphates, 134, 135, 146 ribose, 112 urea, 224, 271 urease, 250, 265, 270, 271 xanthosinate 5⬘-monophosphate, 146 Acireductone, 477, 478, 481, 488–492 Acireductone dioxygenase, 182, 477–481, 486, 528 active site, see Active sites Co(II), 474, 479 enzymatic studies, 489 Fe(II), 474, 479, 480, 483, 486, 489–491, 496 homologs, 481–483 Klebsiella oxytoca, 474–494 mechanistic considerations, 490–493 Mg(II), 474, 479 Mn(II), 474, 479 model, 229 mutation, 475, 480, 490 nickel in, 473–496 ‘one protein, two enzymes’, 477–481, 486 related enzymes, 493–495 sequence identity, 482 spectroscopy, see individual methods structures, 483–486 wild-type, 479, 480, 490 A-cluster, see Clusters Aconitase, 553 ACS, see Acetyl-coenzyme A synthases ACS/CODH, see Acetyl-coenzyme A synthases/carbon monoxide dehydrogenases Actinobacillus pleuropneumoniae, 560 Active sites of (see also Cluster) acetyl-coenzyme A synthase, 203, 205, 207, 209, 214, 522, 523 acireductone dioxygenase, 483, 486–490, 493–495, 522, 523, 528 alcohol dehydrogenase, 272 carbon monoxide dehydrogenase, 200, 202, 522–524 [Fe]-only hydrogenase, 192, 197

SUBJECT INDEX

[Active sites of] glyoxalase I, 455–458, 462, 522, 523 methyl-coenzyme M reductase, 337–340, 344, 347, 349, 350, 522, 523 nickel enzymes, 181–230 nickel superoxide dismutase, 426, 428, 429, 434, 436, 437, 522, 523, 526 [NiFe] hydrogenase, 192, 193, 195, 212, 282, 284, 286, 289, 290, 292, 293, 295, 300, 301, 303, 305, 306, 308, 311, 522, 523 urease, 214–229, 244, 245, 249–266, 268, 269, 272, 522, 523, 555 Adefovir, see 9-[2-(Phosphonomethoxy)ethyl]adenine Adenine(s) (and residues), 141, 167, 168 8-aza-, 146 9-methyl-, 138 9-methyl-1,3-dideaza-, 144 Pt(II) complex, 145 structure, 118 Adenosine, 112 1,N6 -etheno-, see 1,N6 Ethenoadenosine 2⬘-deoxy-, 112 acidity constant, 151 ε-, see 1,N6 -Ethenoadenosine methylthio-, see Methylthioadenosine Ni(II) complex, 119, 120 self-association, 139 structure, 118 Adenosine 5⬘-diphosphate, see 5⬘-ADP (N1)-oxide complex, 153, 154 Adenosine monophosphates, see 2⬘-AMP, 3⬘-AMP, and 5⬘-AMP Adenosine 5⬘-monophosphate (N1)-oxide acidity constants, see Acidity constants complexes, 150–154 stability constants, see Stability constants Adenosine 5⬘-O-thiomonophosphate acidity constants, see Acidity constants complexes, 154–156 stability constants, see Stability constants Adenosine 5⬘-triphosphate, see 5⬘-ATP S-Adenosylmethionine, 475, 476, 481, 526, 528

663

Adenylate cyclase, 513 5⬘-ADP, 131, 512 Cu(II) complex, 140, 141 metal ion complexes, 142 Ni(II) complex, 140, 141, 143, 153 Aeropyrum pernix, 509 Aerosols (containing) (see also Atmosphere) nickel, 621, 623, 628, 629, 638 Affinity constants, see Stability constants α-Alanine (and residues) hydroxamic acid, see Hydroxamic acid Ni(II) complex, 70 stability constants, see Stability constants β -Alanine (and residues) hydroxamic acid, see Hydroxamic acid Ni(II) complex, 66, 70 stability constants, see Stability constants Albumin Cu(II) complex, 65, 84 human serum, see Human serum albumin -like complex, 83, 84 metal-binding motif, 80 Alcaligenes eutrophus, 560 Alcohol dehydrogenase active site, see Active sites zinc in, 272 Aldolases, 477 Algae (see also individual names), 47 green, 39, 44, 45, 314 urease, 242 Alkali metal ions, see individual names Alkaline earth ions, see individual names Alkaliphilus metalliredigenes, 390 Allergy nickel-induced, 84, 582, 584, 592, 601, 621, 622 skin, see Dermatitis and Skin Alloys containing nickel, 6 Alps nickel deposition, 15, 16 Aluminum(III) hydroxide, see Hydroxides oxide, see Oxides

664

Alyssum, 48 bertolonii, 49, 51, 52 lesbiacum, 50 pintodasilvae, 12 murale, 49 Amides (see also individual names), 80, 81 deprotonated, 66 linkage, 86 oxal-, 65 thio-, 79 Amine(s) (see also individual names) glycosyl-, see Glycosylamine tris(2-aminoethyl)-, 119, 121 Amino acids (see also individual names) acidity constants, 67 Cu(II) complexes, 95 derivatives, 71–76 list of, 67 Ni(II) complexes, 63–97 stability constants, see Stability constants stereoselectivity of nickel complexes, 71 Zn(II) complexes, 95 Aminohydroxamates Ni(II) complexes, 71–74 α-Amino-iso-butyric acid, 87, 88 Ni(II) complex, 77 Ni(III) complex, 93 δ -Aminolevulinic acid 13 C-, 328 2-Amino-6-oxopurine, see Guanine Aminophosphonates Cu(II) complexes, 72 Ni(II) complexes, 71, 72 6-Aminopurine, see Adenine Ammonia, 216, 224–227, 242, 271, 554, 556–558 Amoxicillin, 549 2⬘-AMP metal complexes, 144–146, 164 3⬘-AMP metal complexes, 144–146, 164 5⬘-AMP (complexes of), 131–134 Cu(II), 137, 141

SUBJECT INDEX

[5⬘-AMP (complexes of)] 7-deaza-, see Tubercidin 5⬘monophosphate metal ion complexes, 138 Mg(II), 136, 137, 141, 157 Mn(II), 137, 141 Ni(II), 113, 136, 137, 139, 141, 156 Zn(II), 137 Anabaena cylindrica, 38, 39 [14]aneN4, see Cyclam Angiogenesis, 85 Angiotensin II, 85 Animals, see individual species Animal studies of (see also individual species) nickel toxicity, 625–631 Anserine, 90 Antagonistic action, see Interdependencies Antarctica nickel deposition, 15 Anthropogenic (see also Environment) nickel, 2, 7, 14, 15, 41, 42 Antibiotics (see also individual names), 79, 464 gastritis, 547 resistance, 568 treatment of Helicobacter pylori infection, 547–549, 568 Antigens for nickel allergy, 84 Antimicrobial therapy, 547 Antimonide nickel, 4 Antioxidant(s) (see also individual names), 90, 588, 589, 599, 600, 636, 637 metabolism, see Metabolism Apoptosis, 448, 476, 551, 584, 592, 601, 602, 604–607, 644 Arabidopsis thaliana, 47, 51 Archaea (see also individual names), 37 acetyl-coenzyme A synthase/carbon monoxide dehydrogenase, 386–389 carbon monoxide dehydrogenase, 371 corphin coenzyme F430, see F430 corrinoid iron-sulfur proteins, 382, 383

SUBJECT INDEX

[Archaea] methanogenic, 203, 323–350, 384, 524–526 NikR, 562 Archaeoglobus fulgidus, 369–371, 383, 384, 389 Arctic Canadian, 8, 16 nickel concentrations, 2, 8, 16 Norwegian, 2 Arginine (and residues), 84 Ni(II) complex, 68, 83 Arsenide nickel, 4 Arteriosclerosis, 629 Ascorbate (or ascorbic acid), 584, 593–595, 600, 636, 637, 646 oxidation, 593 peroxidase, 44 Asparagine (and residues) Ni(II) complex, 69 Aspartic acid (and residues) Ni(II) complex, 69, 78 Aspergillus niger, 449, 451 japonicus, 493, 494 Association constants, see Stability constants Asthma nickel-induced, 582, 584, 623, 624 Atmosphere (see also Aerosols) nickel in, 2, 6–15, 17, 20 Atomic absorption spectroscopy nickel samples, 4, 5 5⬘-ATP, 475, 512, 607, 631, 637 acidity constants, see Acidity constants -dependent molecular chaperone, 524 dephosphorylation, 161 exchange reaction, 523 hydrolysis, 144, 286, 529, 538, 539 metal ion complexes, 141–143, 164, 165 phosphohydrolase, 369 stability constants, see Stability constants structure, 130

665

ATPase(s), 534, 539, 560 inhibition, 45 Australia nickel mining, 4 Avena sativa, 19 8-Azaadenine, 146 Azide as inhibitor, 366, 432

B B. japonicum, see Bradyrhizobium pasteurii, see Bacillus subtilis, see Bacillus Bacillus pasteurii, 244–267, 530–532, 536, 555 subtilis, 477, 479, 494, 495 Bacteria(l) (see also individual names), 19, 477 acetogenic, 203, 524 anaerobic, 37 Archaea, see Archaea cyano-, see Cyanobacteria gram-negative, 450, 453, 553, 562 gram-positive, 449, 553 methanogenic, 37, 281 N2-fixing, 38 pathogens, 49 photosynthetic, 391 proteo-, 391, 392 Bacteriochlorin iso-, 187 Bacteriophage ΦX174, 513, 514, 533 BAL, see 2,3-Dimercaptopropanol Barium(II) (complex with) carbonyl binding, 123 cytidine, 126 flavin mononucleotide, 148 pyrimidine, 122 xanthosine 5⬘-monophosphate, 148 Bark as biomonitor, 10 Battery nickel/cadmium, 6 B-cluster, see Clusters

666

Beans Jack, see Jack bean nickel in, 19 soy, see Soybean Berkheya coddii, 49, 51, 52 Bicine acidity constants, 162 stability constants of metal ion complexes, 163 structure, 162 ternary complexes, 161–163 Binding constants, see Stability constants Bioavailability of nickel, 17–20, 46 Biogeochemistry of nickel, 1–21 Biomonitors for nickel, 8–10, 17 bark, 10 lichen, see Lichen moss, see Moss Biosynthesis acetyl-coenzyme A, 380, 381, 384, 385, 387, 388, 390, 392 acetyl-coenzyme A synthases, 359, 360 cobalamin, 386, 527, 534 F430, 526 glutathione, 51 histidine, 50 hydrogenases, 514, 529 nickel superoxide dismutase, 529 [NiFe] hydrogenases, 284–286, 291, 537, 539 phospholipid, 590 polyamine, 476, 477, 528 urease, 563, 564 2,2⬘-Bipyridine, 132, 205 Ni(II) complex, 168 Ni(III) complex, 92, 141 ternary complexes, 164 2-[Bis(2-hydroxyethyl)amino]2(hydroxymethyl)-1,3-propanediol, see Bistris Bis(hydroxyethyl)glycine, see Bicine Bis(imidazol-2-yl)methylamine, 79, 80 Bis(N,N-diethyldithiocarbamoyl)disulfide, see Disulfiram

SUBJECT INDEX

Bistris acidity constants, 163 stability constants, 163 structure, 162 ternary complexes, 161–163 Biuret reaction, 65, 66 Cu(II) complex, 65 Black Sea methanogenic archaea, 343–345 Bleomycin Ni(III) complex, 92 Bonds disulfide, see Disulfide hydrogen, see Hydrogen bonds Ni(III)–alkyl, 340, 341 Ni–C, 190, 333, 379 peptide, 83 persulfide, see Persulfide prolyl, see Prolyl bond Boric acid (or borate) buffer, 161 inhibition of urease, 255, 256, 267–270 Bradyrhizobium japonicum, 37, 529, 533, 560 Brassica juncea, 451 Brassicaceae nickel hyperaccumulation, 40 Breast cancer, see Cancer British anti-Lewisite, see 2,3Dimercaptopropanol Bronze Age, 6 Brucella sp., 560 Bryophytes (see also individual names) 17 Buffers (see also individual names) acetate, 161 acidity constants, 163 Bicine, see Bicine Bistris, see Bistris borate, 161 imidazole, 161 phosphate, 161 stability constants, 163 ternary complexes, 161–164 Tris, see Tris

SUBJECT INDEX

667

C C. acetobutylicum, see Clostridium botulinum, see Clostridium difficile, see Clostridium difficile, see Clostridium hydrogenoformans, see Carboxythermus tetani, see Clostridium thermoaceticum, see Moorella thermoacetica Cabbage nickel toxicity, 18 Cadmium(II) (in/complexes with) chelator, 47 cytidine, 126 flavin mononucleotide, 148 glyoxalase I, see Glyoxalase I hydrogenase, 524 hyperaccumulation, see Hyperaccumulating plants nucleic acid binding, 167 nucleotides, 144, 148 peptides, 80 pyridines, 123, 125, 127 ternary nucleotide complexes, 164 transporter, 38 uridine, 127, 128 Calcineurin, 603 Calcium(II) (in) channel, 627, 629, 645 channel blocker, 603 flavin mononucleotide complex, 148 glyoxalase I, see Glyoxalase I homeostasis, see Homeostasis interdepency with other metal ions, see Interdependencies intracellular, 84 ionophores, 602, 603 metabolism, see Metabolism xanthosine 5⬘-monophosphate complex, 148 Calmodulin, 505, 603 Calorimetry isothermal, 454, 458, 480

Calvin cycle, 44 Campylobacter pyloridis, see Helicobacter pylori Canada Arctic, see Arctic nickel mining and smelting, 4, 8–10, 14, 17, 41, 42 Canavalia ensiformis (see also Jack bean), 37, 242 Canavalin, 485, 528 Cancer (see also Carcinomas, Tumors and individual names) breast, 496, 597 colorectal, 476 gastric, 547 histopathology, 638, 639 ovarian, 483, 597 respiratory tract, 621, 623, 639 Capillary electrophoresis nickel samples, 5 Carbohydrates (see also Sugars and individual names), 111 Carbon 13 C-label, 206, 341 14 C, 346, 624 global cycle, 524, 525 Ni–C bond, 190, 333, 379 Carbon dioxide, 358, 365 14 C, 346 as energy source, 324–326, 341 fixation, 326 formation, 493, 524, 554, 556, 557 reduction, 324, 326, 384, 386, 387, 524 Carbonic anhydrase, 554, 557 Carbon monoxide (in) 13 C, 206 14 C, 346 as signaling molecule, 496 dehydrogenase, see Carbon monoxide dehydrogenase formation, 203, 474, 478, 479, 481, 483, 490–492, 496, 523 [NiFe] hydrogenase, 192, 281–283, 285, 286, 288, 290, 292, 294, 295, 300–303, 310–313, 523 oxidation, 358–393, 524

668

[Carbon monoxide (in)] sequence aligments, 396–401 toxicity, see Toxicity Carbon monoxide dehydrogenases, 182, 211, 326, 429, 514, 524, 530 acetyl-coenzyme A synthases, see Acetyl-coenzyme A synthases/ carbon monoxide dehydrogenases activation, 369 active site, see Active sites Carboxythermus hydrogenoformans, 361, 365, 524 catalytic cycle, 365 characteristics, 370 C-cluster, see C-cluster clusters other than C, see Clusters evolution, 392, 393 inhibitors, 366 metallochaperones, 534 models, 200–211 molecular chaperones, 538 mutants, 369, 371 nickel insertion, 534 nickel, 36, 37 phylogeny, 369–373 properties, 359 proton transfer, see Proton transfer redox potentials, see Redox potentials Rhodospirillum rubrum, 359, 361, 366, 368, 369, 524 sequence analysis, 369–373 spectroscopic properties, 395 structure, 361, 362 Carbonyl group metal ion binding, 115–117, 195 nickel, see Nickel carbonyl Carboxylate(s) (see also individual names) bridging, 216, 219, 249, 252 Carboxydothermus hydrogenoformans, 36, 200, 202, 359, 369, 370, 372, 373, 376, 390, 524, 525 Carcinogenesis (or carcinogenicity) (of) effect of other carcinogens, 641 Helicobacter pylori, 548 in experimental animals, 639–642 mechanisms, 643–645

SUBJECT INDEX

[Carcinogenesis (or carcinogenicity) (of)] nickel, 82, 169, 582–584, 586, 587, 599, 600, 602, 606, 607, 619–646 species susceptibility, 641 Carcinomas (see also Cancer and Tumors) chorio-, 483 gastric, 548 nickel-induced, 638, 639, 641 Cardiovascular system nickel toxicity, 629 Carnosine, 90 homo-, 90 Cat nickel studies, 628 Catalase, 637 iron, 229 Catalytic cycle of acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 379, 380 carbon monoxide dehydrogenase, 365 F430, 188 glyoxalase I, 452, 465 methyl-coenzyme M reductase, 332, 333, 347–349 [NiFe] hydrogenases, 281, 282, 295, 296, 298, 307, 309–313 Catechol dioxygenase, see Dioxygenases C-cluster, 361, 370, 380, 381, 385–388, 390, 392 assembly mechanism, 368 nickel incorporation, 368, 369 nickel-deficient precursor, 366–368 redox properties, 364–368 spectroscopic properties, 395 structural model, 363 CD, see Circular dichroism 5⬘-CDP (complexes with) Cu(II), 140 Ni(II), 139, 140 Cell death, see Apoptosis malignant transformation, 642 nickel transport, 528–530 nickel-exposed, 585–587, 594, 608, 643 Channels calcium, see Calcium sodium, see Sodium

SUBJECT INDEX

Chaperones, 595 kinds of, 520 metallo-, see Metallochaperones molecular, see Molecular chaperones nickel, see Nickel chaperones Charge transfer ligand-to-metal, 185 Chelating agents (see also individual names) Ni(II) complexes, 79, 80 Chemokines, 584, 592, 601, 604 genes, 591 Chelation therapy nickel toxicity, 624 China nickel pollution, 41 Chlamydomonas sp., 45 reinhardtii, 291 Chloride nickel, see Nickel chloride Chlorobium phaeobacteroides, 392 Chlorophyll, 46 heavy metal-substituted, 43, 44 magnesium substitution, 43, 51 nickel-substituted, 43, 44, 48 Chromatin damage, 635, 636, 643, 644 Chromatography of nickel, 5 Chromosome aberration, 635 damage, 585 α-Chymotrypsin, 511 Circular dichroism glyoxalase I, 456 Ni(II)-peptide complexes, 80, 86, 91 Ni(II)-sugar complexes, 113 nickel superoxide dismutase, 214, 433 Citrate bismuth citrate, 549 Ni(II), 18–20, 47, 50, 266 Ti(III), 375, 377, 378 Cladina sp., 10 Cladonia pleurota, 10 Clarithromycin, 549, 568 Cleavage acetate, 359, 388 dihydrogen, 193,194, 312, 313 DNA, 90, 167, 635, 636

669

[Cleavage] hydrolytic, 217 4-nitrophenylphosphate, 217 picolinamide, 218 protein, 90 RNA, 167 thioether bond, 190 Clostridium sp., 389, 390, 450 acetobutylicum, 291, 390 botulinum, 390 difficile, 370, 371, 373, 390 tetani, 390 thermoaceticum, see Moorella thermoacetica Clusters A-, 374–381, 385, 386, 392, 525 B-, 361, 367, 370, 385, 387, 389, 395 C-, see C-cluster cubane, 200, 204, 205, 367, 374, 377, 378, 524 D-, 361, 365, 367, 370, 371, 385, 387–390, 395 E-, 371, 385 F-, 371, 385 Fe3S3, 200 Fe3S4, 201, 283, 289, 292, 300, 301, 362, 366, 367, 370, 387, 395, 524 Fe4S4, 35, 203–206, 230, 283, 289, 292, 300, 311, 312, 361, 367, 370, 371, 374, 376–379, 381–383, 385, 387, 392, 393, 395, 422, 524, 525, 632 FeS, 200, 281–283, 287 Ni2Fe4S4, 210, 525 NiFe3S4, 200, 202, 205, 524 NiFe4S4, 200 NiFe4S5, 200, 523 5⬘-CMP (complexes with), 133, 135 Ba(II), 122 Cd(II), 126 Co(II), 126 2⬘-deoxy-, 126, 158 Ni(II), 134, 139 Pt(II) complex, 122 Coal nickel in, 21 Cobalamins biosynthesis, see Biosynthesis

670

[Cobalamins] cob(I)alamin, 330, 379, 382, 383, 390 cob(II)alamin, 330 methyl-, see Methylcobalamin Cobalt (different oxidation states) (in) contact dermatitis, see Dermatitis hyperaccumulation, see Hyperaccumulating plants transporters, see Transporters Cobalt(II) (complexes with) acireductone dioxygenase, see Acireductone dioxygenase carbonyl group, 117 cytidine, 125, 126 flavin mononucleotide, 148 glyoxalase I, see Glyoxalase I hydroxyl group, 117 low-spin d7, 92 nucleic acid binding, 165–167 AMP (N1)-oxide, 153 pyridine, 123, 127 SlyD, 512 ternary ATP–buffer complexes, 163 urease, 266, 267 uridine, 127, 128 water exchange rate, 111 xanthosine 5⬘-monophosphate, 148 Cobalt(III) peptide complexes, 93 CODH, see Carbon monoxide dehydrogenases Coenzymes (see also individual names) A, 326 B (see also 7-Thioheptanoylthreoninephosphate), 182, 185 F420, see F420 F430, see F430 M, 336–342, 345, 346, 348, 349, 383 Cofactor F430, see F430 Collagen, 590, 593, 646 Coordination spheres, see Active sites Copper (different oxidation states) contact dermatitis, see Dermatitis smelting, see Smelting

SUBJECT INDEX

Copper(I) (in), 44, 206–208 acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 374 Copper(II) (complexes of), 44, 122 albumin, 65 aminophosphonate, 72 and fibrinopeptides, 84 biuret, 65 carbonyl group, 117 crystal structures, see X-ray crystal structures dihydroxyacetone phosphate, 117 1,N6 -ethenoadenosine nucleotides, 150 flavin mononucleotide, 148 glycerol 1-phosphate, 117 histidine, 95 hormone, see Hormones hydroxyl group, 117 in plants, see Plants in quercetin 2,3-dioxygenase, 493–495 in SlyD, 512 in urease, 267 1-methylcytosine, 122, 123 nucleotide, 137, 138, 140–146, 157, 158 peptides, see Peptides phosphate, 130 purine nucleobases, 121, 146 purine-nucleotide (N1)-oxides, 153, 154 pyridine, 123, 125, 126 ribose binding, 112 ternary ATP–buffer, 163 ternary nucleotide complexes, 164 xanthosine 5⬘-monophosphate, 148 Copper(III) tripeptide complexes, 87, 93 Copper-zinc superoxide dismutase, 418–420, 428, 434, 436, 438, 526 Corphin coenzyme F430, see F430 origin of name, 328 tetrahydro-, 328 Corrinoid iron-sulfur proteins, 360, 375, 379, 382–387, 391–393, 525 protein sequence alignment, 405–408 subunit phylogeny, 383, 384 Corynebacterium sp., 643

SUBJECT INDEX

Crystal structures of (see also X-ray crystal structures) acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 374 carbon monoxide dehydrogenases, 360, 362 F430, 328 glyoxylase I, 455–458, 465 methyl-coenzyme M reductase, 334–338, 342 nickel superoxide dismutase, 425, 429, 430, 526 [NiFe] hydrogenases, 291–294 urease, 250, 253, 255, 257, 258, 260, 262, 264–267 5⬘-CTP, 146 complexes, 129, 141, 143 structure, 131 Cubane cluster, see Cluster Cupins, 483, 485, 493, 494, 528 Cyanate (in) hydrolysis, 226, 227 thio-see Thiocyanate urease, 216, 224–227 Cyanide (in), 37 carbon monoxide dehydrogenase, 366 diazotrophic, 38 Fur protein, 553 micronutrient, see Micronutrients nickel, 32–54 nickel superoxide dismutase, 211, 526, 527 [NiFe] hydrogenase, 192, 199, 281–283, 285, 286, 288, 290, 292, 295, 300, 302, 303, 305, 306, 310, 523 toxicity, see Toxicity Cyanobacteria (see also individual names), 314 diazotrophic, 38 Fur, 553 micronutrients, 37–40 nickel in, 32–54 nickel superoxide dismutase, 211, 526, 527 Cyclam (complex with) Ni(I), 188 Ni(II), 189–191

671

[Cyclam (complex with)] Ni(III), 144 tetramethyl-, 188–191 Cyclic voltammetry of F430, 332 Ni(II) complexes, 213 Ni(II)-Ni(0), 210 Ni(II)-Ni(I), 210, 334 Ni(III) complexes, 92 [NiFe] hydrogenases, 197, 198 Cyclophilins, 503–505, 513 Cynops pyrrhogaster, 640 Cysteine (and residues), 84 bridge, 367, 374, 523 in [NiFe] hydrogenases, 286, 300, 308, 523 Ni(II) complex, 66, 69, 80–82, 95, 96 oxidation, 308 seleno-, see Selenocysteine S-methyl-, 338 β,β -dimethyl-, see Penicillamine Cytidine metal ions complexes, 123, 125, 126 structure, 118 Cytidine 5⬘-diphosphate, see 5⬘-CDP Cytidine 5⬘-monophosphate, see 5⬘-CMP Cytidine 5⬘-triphosphate, see 5⬘-CTP Cytochrome b, 280, 283, 287 Cytochrome(s) c, 280, 287 c3, 283 c3 oxidoreductase, see [NiFe] hydrogenase Cytochromes P450, 448 Cytokine(s), 584, 590–592, 601–604, 608 genes, 591, 592 Cytosine, 168 methylation, 584, 586, 607 structure, 118 Cytotoxicity of methylglyoxal, 448, 449, 527

D D. baculatum, see Desulfomicrobium and Desulfovibrio

672

[D.] desulfuricans, see Desulfovibrio ethenogenes, see Dehalococcoides fructosovorans, see Desulfovibrio gigas, see Desulfovibrio hafniense, see Desulfitobacterium vulgaris, see Desulfovibrio Data bases of proteins PDB, see Protein Data Bank D-cluster, 361, 365, 367, 370, 371, 385, 387–390, 395 Decarboxylases ornithine, 476 oxalate, 493, 494 Deficiency of nickel, 39, 40, 624, 625, 630 nitrogen, 40 Dehalococcoides ethenogenes, 382, 390 Dehydrogenases carbon monoxide, see Carbon monoxide dehydrogenase glucose-6-phosphate, 597 glyceraldehyde-3-phosphate, 448, 597 lactate, 446, 447 methylene tetrahydromethanopterin, 325 Density functional theory calculations F430, 188, 331, 332, 341, 348 nickel superoxide dismutase, 436 [NiFe] hydrogenase, 301, 303–306, 310, 312, 313 urease, 269, 271 2⬘-Deoxyadenosine, 112 1-(2⬘-Deoxy-β -D-ribofuranosyl)thymine, see Thymidine Deoxyribonucleic acid, see DNA 2⬘-Deoxyribose, 112 Deprotonation constants, see Acidity constants Dermatitis (induced by) allergic contact, 582, 592, 624, 630 cobalt, 624 copper, 624 nickel, 621, 623, 624, 629, 637 Deschampsia flexuosa, 18 Desulfitobacterium hafniense, 369, 370, 372, 390 Desulfomicrobium baculatum, 521, 523

SUBJECT INDEX

Desulfovibrio baculatum, 292 desulfuricans, 292, 370, 391 fructosovorans, 199, 289, 290, 292 gigas, 36, 196, 199, 281, 289, 292, 293, 295–299, 305, 421, 521 vulgaris, 289, 292, 294, 295, 297, 298, 301–303, 305, 306, 391 Detoxification (of) (see also Toxicity) chelation therapy, see Chelation therapy mechanism in plants, 46–48 methylglyoxal, see Methylglyoxal nickel, 46, 624 reactive oxygen species, 551, 552 superoxide radicals, 551 DFT calculations, see Density functional theory calculations Diamidophosphate urease complex, 256, 257, 268–272 1,2-Diaminoethane, see Ethylenediamine 1,4-Diaminobutane, see Putrescine 2,4-Diaminobutyric acid hydroxamic acid, see Hydroxamic acid Ni(II) complex, 69, 71 1,3-Diaminopropane Ni(II) complex, 70, 71, 112, 113 2,3-Diaminopropionic acid hydroxamic acid, see Hydroxamic acid Ni(II) complex, 69, 71 2,6-Diaminohexanoate, see Lysine 2,5-Diaminopentanoate, see Ornithine Dichotomy of metal ion binding to purines, 120, 121 Dielectric constant reduced, 117, 149 Dien, see Diethylenetriamine Diethyldithiocarbamate sodium, 624 Diethylenetriamine (complexes of) Ni(III), 92 Pt(II), 122 Diethylenetriamine-N,N,N⬘,N⬙N⬙pentaacetate, 19 prevention of contact dermatitits, 624 Diglycine, see Glycylglycine

SUBJECT INDEX

Dihydrogen (see also Hydrogen) as energy source, 324, 325, 341 cleavage, 193, 194, 312, 313 cycling, 38 dissociation, 280 [NiFe] hydrogenases, 281, 282, 289, 290, 293–295, 311 oxidation, 38, 384, 387, 521 producing catalysts, 191 production, 193, 514 Dihydroxyacetone phosphate (complexes with), 115, 446, 448 Cu(II), 117 Ni(II), 116, 117 2,3-Dimercaptopropanol nickel toxicity, 624 Dioxygen (see also Oxygen) (in), 422, 593, 593 acireductone dioxygenases, 479, 489–491, 493 formation, 418, 526 [NiFe] hydrogenases, 294, 295, 308, 311, 314 oxidation, 495 reduction, 633 Dioxygenases acireductone, see Acireductone dioxygenase catechol, 463 quercetin 2,3-, 493–495 N,N⬘-Disalicylideneethylenediamine, see Salen Diseases (see also individual names) heart, 418 inflammatory (see also Inflammation), 418 intestinal tract, 545–570 ischemic, 418 neurodegenerative, see Neurodegenerative diseases Dismutases superoxide, see Superoxide dismutase Dissociation constants (see also Equilibrium constants and Stability constants) urease, 252, 261

673

Disulfide(s), 422 bonds, 78, 259, 425 bridge, 84, 379 formation, 325, 326, 333, 334 Ni(II) complex, 78, 79 radical, see Radicals redox potential, see Redox potentials Disulfiram, 624 Dithiol (see also Thiols) nickel complex, 333 Dithionite in acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 375, 377 carbon monoxide dehydrogenases, 364 DNA B-, 165 calf thymus, 167 cleavage, 90, 167, 635, 636 damage, 604–606, 608, 635, 636 duplex, 167–169 effects of Ni(II), 89 fragmentation, 605 inhibition of repair enzymes, 644 insertion, 287, 288 M-, 167, 168 methylation, 475, 583, 586, 587, 599, 607, 644 microarray analysis, see Microarray analysis of DNA nickel binding, 632 oxidative damage, 90, 585, 635, 636, 643, 644, 646 –protein cross-linking, 635 recognition, 90 synthesis, see Synthesis synthetic analogs, 80 Z-, 165 Dog nickel studies, 630 Drugs, see individual names dTDP, see Thymidine 5⬘-diphosphate dTMP, see Thymidine 5⬘-monophosphate DTPA, see DiethylenetriamineN,N,N⬘,N⬙N⬙-pentaacetate Duodenum ulcer, 546 dTTP, see Thymidine 5⬘-triphosphate

674

SUBJECT INDEX

E E. coli, see Escherichia Earth’s crust (see also Soil) nickel in, 4, 6, 110 E-cluster, see Clusters EDTA, see Ethylenediamine-N,N,N⬘,N⬘tetraacetate Electrochemistry of [NiFe] hydrogenases, 306–309 Electrodes glassy carbon, 335 graphite, 306, 307, 309 platinum, 309 standard calomel, 334, 335 Electron density [NiFe] hydrogenases, 294, 295, 298, 299 urease, 251, 252, 256, 261, 265 Electronic spectra, see Absorption spectroscopy Electron microprobe analysis nickel samples, 5 Electron nuclear double resonance, see ENDOR Electron paramagnetic resonance, see EPR Electron potentials, see Redox potentials Electron spin echo envelope modulation, see ESEEM Electron spin resonance, see EPR Electron transfer (in) acetyl-coenzyme A synthase, 204 acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 380, 385 carbon monoxide dehydrogenases, 361–363, 365, 373, 388, 390, 391 Ni(III)/Ni(II)-peptide complexes, 88 [NiFe] hydrogenases, 282, 283, 293, 298, 313, 325 proton-coupled, 418–420 superoxide dismutases, 418, 419 Electrophoresis capillary, 5 polyacrylamide gel, 453 SDS-PAGE, see Sodium dodecylsulfate polyacrylamide gel electrophoresis

Electrospray ionization mass spectrometry acetyl-coenzyme A synthase models, 209 nickel superoxide dismutase, 428 Elodea, 41 Empetrum nigrum, 18 ENDOR spectroscopy (of) 1 H, 304, 305 17 O, 305 C-cluster of carbon monoxide dehydrogenase, 364, 365 F430, 330, 331, 341, 342 Ni(I) complexes, 187 [NiFe] hydrogenases, 192, 193, 281, 282, 303–305 Enterobacteriaceae, 446 Environment nickel in, 1–21, 32–54 nickel pollution, 40–48, 524 Environmental Protection Agency of the United States nickel concentrations, 4, 5 Enzymes (see also Proteins and individual names) models of active sites, 181–230 EPR (studies of) acetyl-coenzyme A synthase, 204, 206 F430, 183, 330–332, 338 glyoxalase I, 457 methyl-coenzyme M reductase, 335, 338–342 Ni(I) complexes, 186, 187, 189, 210, 377, 380 Ni(III) complexes, 65, 89, 91–93, 193, 380 nickel enzymes, 34 nickel superoxide dismutase, 214, 429, 431, 432 [NiFe] hydrogenases, 192, 193, 281, 290, 291, 298, 300–306 Equilibrium constants (see also Acidity constants and Stability constants), 121, 126, 153 definition, 132, 133 intramolecular, 125, 132 methyl-coenzyme M reductase, 346 Erythropoietin, 629, 640 nickel-induced expression, 592, 593

SUBJECT INDEX

Escherichia coli, 528 acetyl-coenzyme A synthase/carbon monoxide dehydrogenase, 373, 376 acireductone dioxygenase, 474, 479, 491, 493 carbon monoxide dehydrogenase, 366, 367, 369 Fur, 553 glyoxalase I, 451–465 methylglyoxal, 446, 449, 450 nickel mutagenicity, 643 nickel superoxide dismutase, 39, 423, 425, 527 [NiFe] hydrogenase, 285, 286, 327, 523, 532, 533, 537 NikR, 562, 563 peptidyl prolyl cis/trans isomerases, 504, 505 SlyD, 502, 504–510, 513, 514, 533 ESEEM four-pulse, see Hyperfine sublevel correlation spectroscopy ESI-MS, see Electrospray ionization mass spectrometry ESR, see EPR 1,N6 -Ethenoadenosine acidity constants, see Acidity constants nucleotide complexes, 150, 151 stability constants, see Stability constants structure, 151 N,N⬘-Ethylenebis(salicylideneaminato), see Salen Ethylenediamine (complex with), 208 Ni(II), 70, 71, 112–114 Ni(III), 92 Ethylenediamine-N,N,N⬘,N⬘-tetraacetate (complex with), 451, 474 Ni(II), 18, 19, 41, 266, 567 Eukaryotes (or eukaryotic) (see also individual names), 477, 481, 483 DNA, 586 FK506-binding proteins, 503, 509 glyoxalases, 446, 450, 451, 465 [NiFe] hydrogenases, 287 Evolution of acetyl-coenzyme A decarbonylase/ synthases, 386–393

675

[Evolution of] carbon monoxide dehydrogenases, 392, 393 metalloenzymes, 373, 392, 393 EXAFS studies of acetyl-coenzyme A synthase, 204 acireductone dioxygenase, 474, 480 glyoxalase I, 455 methyl-coenzyme M reductase, 339, 343 Ni(I) complexes, 187 nickel superoxide dismutase, 426, 427 [NiFe] hydrogenases, 192, 299, 300 urease, 258 Excluders, 33, 48 Extended absorption fine structure spectroscopy, see EXAFS

F F420 -reducing [NiFe] hydrogenase, 325, 387, 388 F430 (see also Methyl-coenzyme M reductase), 35, 37, 187, 323–350, 523, 525, 529 biosynthetic pathway, 526 catalytic cycle, 188 ESR studies, see EPR magnetic circular dichroism studies, see Magnetic circular dichroism methyl-, 332, 333 models, 182–191 Ni(I), see Nickel(I) Ni(III), see Nickel(III) nonplanarity, 329 properties, 330–333 redox potential, see Redox potentials structure, 183, 325, 328–335 thiol coordination, 341, 342 Fabaceae, 19 Familial amyotrophic lateral sclerosis, 418 F-cluster, see Clusters FeMo cofactor formation, 369 Fenton-like reaction, 89, 551 [Fe]-only hydrogenases, 191, 192, 291, 521 active site, see Active sites

676

SUBJECT INDEX

Ferredoxins (see also individual names) [4Fe4S], 35, 422 reduction, 326, 380 Ferritins, 553, 554, 589, 627 Fertilizers containing nickel, 2, 3 nitrogen, 242 Fibrinogen, 84 Finland nickel pollution, 2, 7–15, 18, 20, 41 Flavin mononucleotide acidity constants, see Acidity constants complexes, 148, 149 stability constants, see Stability constants structure, 147 FMN, see Flavin mononucleotide Formaldehyde, 375 Formate, 325 production, 388 Formation constants, see Equilibrium constants and Stability constants Fosfomycin, 464 resistance protein, 463, 464 Fourier transform infrared spectroscopy (studies of) [NiFe] hydrogenases, 281, 282, 290, 292, 295–299 stopped flow, 295 Fructose Ni(II) complex, 113 FTIR, see Fourier transform infrared spectroscopy Fulvic acid nickel complex, 4, 19 Fungi (or fungal) ectomycorrhizal, 46 glyoxalases, 451 pathogens, 49 urease, 242

G Gastritis, 35, 346 Helicobacter pylori-induced (see also Helicobacter pylori), 547, 548

Gastrointestinal tract diseases, see Diseases infection, 242 5⬘-GDP complexes of Cu(II), 140, 142 metal ions, 142 Ni(II), 140, 141, 143 Gel electrophoresis polyacrylamide, 453 Genes acetyl-coenzyme A synthases/ carbon monoxide dehydrogenases, 389–392 coding for cytokines and chemokines, 591, 592 coding for extracellular matrix proteins, 590, 591 coding for metal binding, 588, 589 nickel-dependent expression, 581–609 stress response, 605 tumor-suppressor, 606, 607 Genome sequences of acetyl-coenzyme A decarbonylase/synthases, 409, 410 Geobacter metallireducens, 391 sulfurreducens, 391 Glucose (see also Sugars) 6-phosphate dehydrogenase, see Dehydrogenases transport, 597 Glutamic acid (or glutamate) hydroxamic acid, see Hydroxamic acid Ni(II) complex, 68, 78 Glutamine Ni(II) complex, 68 Glutathione, 47, 447, 464, 527, 584, 600, 628, 631, 636, 637 air oxidation, 82 biosynthesis, see Biosynthesis derivatives, 458–460 metabolism, see Metabolism Ni(II) complex, 78, 81, 82, 632 peroxidase, see Peroxidases reduced, 90 reductase, see Reductases S-lactoyl-, 449, 450, 455, 458, 527

SUBJECT INDEX

[Glutathione] S-transferase, see Transferases synthetase, see Synthases Glyceraldehyde-3-phosphate, 447, 448 dehydrogenase, see Dehydrogenases Glycerol 1-phosphate acidity constant, 117 Cu(II) complex, 117 metal ion complexes, 149 Ni(II) complex, 115–117 structure, 147 Glycine α-mercaptopropionyl-, 82 di-, see Glycylglycine γ -glutamylcysteinyl-, see Glutathione hydroxamic acid, see Hydroxamic acid Ni(II) complex, 68, 69, 71 N-methyl-, see Sarcosine oligo-, 76 tetra-, see Tetraglycine thio-, 338 tri-, see Triglycine Glycine max. (see also Soybean) nickel uptake, 18 Glycosides N-, see Glycosylamines Glycosylamines Ni(II) complexes, 112, 113 Glycylglycine Ni(II) complex, 76, 93 Ni(III) complex, 93 Glycylglycylglycine, see Triglycine Glycylglycylglycylglycine, see Tetraglycine Glyoxal methyl-, see Methylglyoxal phenyl-, 446, 451 Glyoxalase I, 37, 527, 528 active site, see Active sites Ca(II), 462 catalytic cycle, 452, 465 catalyzed isomerization, 458–460 Cd(II), 453, 454, 456, 458, 462, 527 classes, 452 Co(II), 453–456, 458, 460, 462, 463, 527 crystal structures, see Crystal structures

677

[Glyoxalase I] Escherichia coli, see Escherichia coli genes, 460–463 Mg(II) in, 451, 452, 454, 462 mechanistic studies, 453–460 member of the βαβββ superfamily, 463, 464 metal activation, 451, 452, 460–463 Mn(II), 453–455, 457, 462, 527 mutagenesis studies, 462 nickel-dependent, 445–466 sequence alignments, 461 various sources, 450, 451 Zn(II), 451, 452, 454–463, 465, 527 Glyoxalase II, 446, 447, 450, 451, 527 2⬘-GMP, 145 2⬘d3⬘-, 145 3⬘-GMP, 145 3⬘,5⬘-c, 114 5⬘-GMP (complexes with), 133 Cu(II), 137 d-, 113, 114, 158, 164 Mg(II), 136, 137 Mn(II), 137 Ni(II), 113, 114, 135–139 stability constants, 137 Zn(II), 137 Goat nickel studies, 625 Greenland ice core, 15 nickel accumulation, 15 Growth factors transforming, 588, 590 5⬘-GTP, 37, 512 binding motifs, 533 -dependent molecular chaperones, see Molecular chaperones -dependent urease activation, 536 hydrolysis, 536, 538, 539, 556 metal ion complexes, 143 Ni(II) complex, 141, 143 GTPases, 533, 536, 538, 559 Guanine (and moiety), 141, 165–167 acidification, 122 9-methyl-, 138 8-oxo-, 636

678

SUBJECT INDEX

[Guanine (and moiety)] Pt(II) complex, 145 structure, 118 Guanosine 2⬘-deoxy-, 165 8-hydroxy-2⬘-deoxy-, see 8-Hydroxy2⬘-deoxyguanosine Ni(II) complex, 121, 122 self-association, 139 structure, 118 Guanosine 5⬘-diphosphate, see 5⬘-GDP Guanosine monophosphate, see GMP Guanosine 5⬘-triphosphate, see 5⬘-GTP Guinea pig nickel studies, 592, 593, 629, 630, 637

H H. felis, see Helicobacter felis influenzae, see Haemophilus pylori, see Helicobacter pylori Haber-Weiss-Fenton reaction, 551 Haemophilus influenzae, 505 Haldane equation, 346 Hamster nickel studies, 629, 641 Helicobacter felis, 569 Helicobacter pylori, 35, 534 antibiotic treatment of infection, 547–549, 568 associated diseases, 548 chemotaxis, 550 diagnostics, 549 drug targets, 568, 569 Fur, 564–566 genes, 555, 559, 562–567, 569 genome, 547, 562, 565 hydrogenases, 559, 560, 565 -induced gastritis, 547 infection, 548, 549 mechanism of pathogenesis, 549–551 metabolism, see Metabolism metal ion export, 567, 568 microbiology, 547 nickel transporters, 560, 561, 566

[Helicobacter pylori] NikR, 562–566, 570 role of nickel, 545–570 urease, 242–267, 521, 529–532, 536, 537, 550–559, 564 Heme oxygenase, 589 Hepatitis B, 156 C, 481, 483 Herbivores, 40, 49, 51 nickel concentrations, 12 Heteronuclear single quantum correlation 1 15 H, N-, 496 Hexokinase I, 597 High-potential iron proteins, 422 Histamine complex with Ni(II), 70, 71 Ni(III), 94 Histidine (and residues) (see also Imidazole) (complexes with) Cu(II), 95 fragile triad, 607 hydroxamic acid, see Hydroxamic acid kinase, see Kinases Ni(II), 66, 69, 71, 80–83, 95, 96 Ni(III), 94 nickel binding, 50, 51 N-methyl-, 338 ternary complexes, 164 Zn(II), 95 Histone(s) acetylation, 587, 605, 608, 644, 645 methylation, 584 modification, 583, 587 Ni(II) complexes, 82–86, 90, 632, 633, 635, 636 transcription, 587 Homeostasis of calcium, 602–604, 645 iron, 562, 632 metal ions, 552 methylthioadenosine, 477 nickel, 514, 562 Homo sapiens, see Human Hormones (complexes with) (see also individual names) Cu(II), 86

SUBJECT INDEX

[Hormones (complexes with)] gonadotropin-releasing, 86 hypothalamic, 86 luteinizing-hormone releasing hormone, 86 neuro-, 85 Ni(II), 86 peptides, see Peptides Zn(II), 86 HSQC, see Heteronuclear single quantum correlation Human fibrinopeptide A, 84 metal diet, 569 protamine, 84 sperm, 84 Human serum albumin, 84 Ni(II) binding, 623 Humic acid nickel complex, 4, 19 Hydride bridge, 305, 311–313 Hydrogen (see also Dihydrogen) 1 H ENDOR, see ENDOR cycle, 38 formation, 308, 314, 331, 390 gas as energy carrier, 39 metabolism, see Metabolism Hydrogenases, 514, 521–524, 530, 537, 538 biosynthesis, see Biosynthesis catalytic, 304–306, 310 classes, 280 cytosolic, 284, 290 de-, see Dehydrogenases [FeFe], 280, 314 [Fe]-only, see [Fe]-only hydrogenase Fe-S cluster-free, 191, 325, 521 inhibition, 569 membrane-bound, 280, 284, 288, 309 metallochaperones, 532, 533 molecular chaperones, 537, 538 nickel insertion, 369 nickel-iron, see [NiFe] hydrogenases [NiFeSe], 291, 292, 301 periplasmic, 284 regulatory, 280, 304, 306 soluble, 280, 284, 289, 295, 300

679

Hydrogen bond (in/to) acireductone dioxygenase, 485 carbonyl group, 114, 138, 139, 165, 166 hydroxyl group, 163, 420 Ni(II) complexes, 113, 114 [NiFe] hydrogenases, 306 Pt(II) complexes, 159 ribose, 112 superoxide dismutases, 420, 434, 435, 437, 438 urease, 215, 219, 220, 251–259, 262, 265, 266, 269–272 Hydrogen peroxide (see also Peroxides), 212, 214, 418, 419, 422, 427, 428, 436–438, 551, 602, 604, 633–636, 644, 646 activation, 89, 90 disproportionation, 90 formation, 493 Hydrolases (see also individual names) S-2-hydroxyacylglutathione, see Glyoxalase II phospho-, 369 Hydrolysis 5⬘-ATP, 144, 286, 529, 538, 539 cyanate, 226, 227 5⬘-GTP, 536, 538, 539, 556 peptide bond, 83 phenylphosphorodiamidate, 268, 270 urea, 226, 243, 249, 256, 265, 267, 268, 554, 556 Hydroperoxide (see also Peroxides) in [NiFe] hydrogenases, 294, 306, 308, 310, 312 Hydroxamates (or hydroxamic acids) (see also individual names) α-alanine-, 73, 75 β -alanine-, 73, 74 aceto-, see Acetohydroxamic acid acidity constants, 72, 73 amino-, see Aminohydroxamic acid bridge, 259 2,3-diaminopropiono-, 74 2,4-diaminobutyric acid, 74 glutamic acid-γ -, 74 glycine-, 73 histidine-, 74 sarcosine, 73

680

SUBJECT INDEX

Hydroxide(s) aluminum, 13 bridging nickel and iron, 367 bridging two nickel, 217, 218, 220, 221, 225, 252, 255, 258, 262, 265, 269–272, 305, 306, 310, 311 iron, 13 Hydroxo complexes, 111 S-2-Hydroxyacylglutathione hydrolase, see Glyoxalase II 8-Hydroxy-2⬘-deoxyguanosine, 90 6-Hydroxydopamine, 418 Hydroxyl groups (see also Sugars) bridge in carbon monoxide dehydrogenase, 364 metal ion coordination, 112–117 radicals, see Radicals ribose, see Ribose Hydroxylases (see also individual names), 593–598 Hylocomium splendens, 9 Hyperaccumulating plants (see also individual names), 33, 38, 46 cadmium, 50 cobalt, 52 mechanisms, 50, 51 nickel, 12, 17, 21, 40, 45, 47–53 zinc, 48 Hyperfine sublevel correlation spectroscopy methyl-coenzyme M reductase, 340, 342 [NiFe] hydrogenases, 193, 304, 306 Hyperglycemia, 449 Hypogymnia physoides, 9 Hypoxanthine, 114, 118, 121, 141 acidification, 122 Hypoxia, 583, 584, 587, 590, 593, 595–598, 608, 636, 640 HYSCORE, see Hyperfine sublevel correlation spectroscopy

I Iceland nickel deposition, 8

ICP-AES, see Inductively coupled plasma-atomic emission spectrometry ICP-MS, see Inductively coupled plasmamass spectrometry ICP-OES, see Inductively coupld plasmaoptical emission spectrometry 5⬘-IDP (complexes with) Cu(II), 140, 142 metal ions, 142 Ni(II), 140–142 (N1)-oxide, 154 Imidazole (and moieties) (see also Histidine), 332 bridge, 420 in ternary complexes, 164 1-methyl-, 120 1-methyl-4-aminobenz-, 144 1-methylbenz-, 119 Ni(II) complexes, 69, 80 Immune system nickel attack, 624, 629, 630 Implants (see also Prostheses) surgical, 592, 621, 624 5⬘-IMP (complexes with), 131, 133 metal ions, 137 Mg(II), 136, 137 Ni(II), 113, 114, 135–139 (N1)-oxide, see Inosine 5⬘monophosphate (N1)-oxide Indonesia nickel mining, 4 Inductively coupled-plasma atomic emission spectrometry glyoxalase I, 458 Inductively coupled plasma-mass spectrometry nickel samples, 4–6 Inductively coupled plasma-optical emission spectrometry nickel samples, 4 Inflammation (or inflammatory) gastric, 548–552 nickel-induced, 600, 621, 624 respiratory system, 582, 589 response, 477, 584, 550, 601, 604, 608, 646

SUBJECT INDEX

Infrared (spectroscopy) studies (of) acetyl-coenzyme A synthase models, 204, 206, 208, 210 Fourier transform, see Fourier transform infrared spectroscopy [NiFe] hydrogenase (models), 192, 199, 297 Inhalation of nickel carbonyl, 582, 583, 621–623, 626–629 nickel-containing dust, 621 nickel oxide, 628, 638, 639 nickel subsulfide, 628, 638–641 nickel sulfate aerosol, 628, 629, 640 Inhibition of adenylate cyclase, 513 ATPases, 45 plant growth, 33, 42–45 hydrogenases, 569 methanogenesis, 388 methyl-coenzyme M reductase, 331, 345, 349 nickel superoxide dismutase, 432 [NiFe] hydrogenase, 294, 308 peptidyl prolyl cis/trans isomerases, 515 photosynthesis, 43–45, 51 root function in plants, 42–44 SlyD, 511, 512 urease, 228, 255–263, 267–272, 569 Inosine Ni(II) complex, 120–122 self-association, 139 structure, 118 Inosine 5⬘-diphosphate, see 5⬘-IDP Inosine 5⬘-monophosphate, see 5⬘-IMP Inosine 5⬘-monophosphate (N1)-oxide complexes, 150–154 structure, 151 Inosine 5⬘-triphosphate, see 5⬘-ITP Interdependencies (between metal ions) calcium–nickel, 642 iron–nickel, 642 magnesium–nickel, 43, 48, 642 manganese–nickel, 642 nickel–zinc, 642 Ionization constants, see Acidity constants

681

Intestinal tract, see Gastrointestinal tract Interleukin-1, 592, 598, 600, 601, 604 Ionophores (see also individual names) A23187, 602 ferric, see Siderophores ionomycin, 603 IR, see Infrared (spectroscopy) studies (of) Iron (different oxidation states) (in) 57 Fe, 206, 303 carbonyls, 195 channel, 627 homeostasis, see Homeostasis hydroxide, see Hydroxide(s) interdependency with other metal ions, see Interdependencies metabolism, see Metabolism oxide, see Oxides transport, 38 uptake regulator Fur, 553, 564, 565 Iron(0), 195, 642 Iron(II) in acireductone dioxygenase, see Acireductone dioxygenase [NiFe] hydrogenases, see [NiFe] hydrogenases quercetin 2,3-dioxygenase, 493–495 Iron(III) (in), 642 superoxide dismutases, 421 Iron-molybdenum cofactor, 369 Iron regulatory proteins, 632 Iron-sulfur proteins (see also individual names) corrinoid, see Corrinoid iron-sulfur proteins Iron superoxide dismutase, 418–421, 423, 428, 434, 438, 526 Irving-Williams series, 96, 129, 138 Isomerases SlyD, see Peptidyl prolyl cis/trans isomerase SlyD triose phosphate, 447, 448, 460 Isomeric complex equilibria, 115, 117, 132, 137, 142, 143, 145, 153, 156–158, 168 Isotope labeling (see also individual elements), 301

682

SUBJECT INDEX

5⬘-ITP in ternary complexes, 164 metal ion complexes, 143 Ni(II) complex, 141, 143, 164

J Jack bean canavalin, 485, 528 urease, see Urease

K Kidney nickel toxicity, 629 stone formation, 242 Kinases histidine, 284 methylthioribose, 477 phosphatidylinositol-3-, 604 Klebsiella aerogenes, 37, 244–267, 521, 530, 531, 535, 536, 556 oxytoca, 474–481, 528 Kola Peninsula nickel deposition, 8–11, 14, 18, 20, 42 Kupfernickel, 6, 34

L Lactate D-, 446, 447, 450 dehydrogenase, see Dehydrogenases production, 446, 447 Lactotransferrin, 588, 589 Lakes (see also Water) nickel in, 10, 11, 14, 41 Legumes (see also individual names), 19 N2-fixing, 38 Leishmania major, 464, 465 Lemna, 41, 45 Leucine ternary complexes, 164

Lichens as biomonitors for nickel, 9, 10, 17 Ligand–ligand interactions, 160, 165 Lignin, 45 Lipid metabolism, see Metabolism peroxidation, 43, 44, 600, 628, 629, 633, 634 Liver damage, 628 nickel toxicity, 628 Loffler⬘s syndrome nickel-induced, 623, 624 Lung fibrosis, 623, 642 inflammation, 628, 641 injury, 583, 587–592 Lymphomas, 547, 548 Lysine hydroxylation, 593 Ni(II) complex, 593

M M. acetivorans, see Methanosarcina barkeri, see Methanosarcina burtonii, see Methanococcoides hungatii, see Methanospirillum jannaschii, see Methanococcus and Methanocaldococcus kandleri, see Methanopyrus marburgensis, see Methanothermobacter maripaludis, see Methanococcus mazei, see Methanosarcina stadtmanae, see Methanosphaera thermoacetica, see Moorella thermoautotrophicum, see Methanobacterium thermoautotrophicus, see Methanothermobacter thermolithotrophicus, see Methanococcus thermophila, see Methanosarcina thermophilus, see Methanoculleus

SUBJECT INDEX

[M.] tuberculosis, see Mycobacterium urea, see Micrococcus vaniellii, see Methanococcus voltae, see Methanococcus Macrochelates ATP, 143, 164 intramolecular, 142, 143, 159 Ni(II)-peptides, 78 nucleoside 5⬘-monophosphates, 137, 138, 144–150, 155, 164 nucleoside 5⬘-triphosphates, 143 purine-nucleoside 5⬘-phosphates, 131, 132, 137, 141 purine-nucleoside 5⬘-diphosphates, 142 Macroconstants, see Acidity constants and Stability constants Macrocycles (see also individual names) (complexes with), 34, 35, 187 conformational change, 186 cyclam, 188 F430, 329–331, 334 {N4}, 183, 186 Ni(II), 79 Macrophages, 591, 597, 630 Magnesium(II) (in/complexes with), 122 acireductone dioxygenase, see Acireductone dioxygenase cytidine, 123, 125 electron transport, 282 flavin mononucleotide, 148 glyoxalase I, see Glyoxalase I interdependency with other metal ions, see Interdependencies [NiFe] hydrogenases, 282–284, 289, 292, 293 nucleotides, 135, 142, 144, 148, 157 phosphate, 130 pyridine, 123, 125 thiophosphate, 155 transporter, 38 Magnetic circular dichroism (studies of) acetyl-coenzyme A synthase, 378 carbon monoxide dehydrogenase, 395 F430, 183 methyl-coenzyme M reductase, 343

683

[Magnetic circular dichroism (studies of)] nickel superoxide dismutase, 429, 432–434 soft X-ray, 378 urease, 258 Magnetic susceptibility measurement of urease, 258 Malate, 47 MALDI-MS, see Matrix-assisted UV laser desorption/ionization mass spectroscopy MALDI-TOF mass spectrometry F430, 327, 329, 344 Maltose Ni(II) complex, 113 Mandelate D-, 446 production, 446 Manganese (different oxidation states) interdependency with other metal ions, see Interdependencies oxide, see Oxides Manganese(II) (in/complexes with) acireductone dioxygenase, see Acireductone dioxygenase carbonyl group, 117 glyoxalase I, see Glyoxalase I hydroxyl group, 117 nucleotides, 137, 142, 144, 157 oxalate decarboxylase, 493, 494 oxalate oxidase, 493, 494 phosphate, 129 purine-nucleotide (N1)-oxides, 148, 153 pyridine, 127 ternary nucleotide complexes, 164 uridine, 128 Manganese(III), 144 Manganese superoxide dismutase, 418–421, 434, 438, 526 Matrix-assisted laser desorption ionization time-of-flight, see MALDI-TOF mass spectrometry Matrix metalloproteinases, 483 MCD, see Magnetic circular dichroism Melanostatin, 85, 86 β -Mercaptoethanol, 258, 259

684

7-Mercaptoheptanoylthreonine, see Coenzyme B 3-Mercaptopropionic acid, 208 Mesopotamia, 6 Metabolism (of) (see also Homeostasis) antioxidants, 584 calcium, 602 glutathione, 600 Helicobacter pylori, 547 hydrogen, 38, 284, 287 iron, 564, 589, 595 lipid, 42 methylglyoxal, 449 methylthioadenosine, 479 nickel, 519–539, 563, 566–568 nitrogen, 555 plant, 32–54 Metal fume fever nickel-induced, 623, 624 Metal ions, see individual names Metallochaperones, 520, 521, 529–534, 538 carbon monoxide dehydrogenase, see Carbon monoxide dehydrogenases copper, 531 mechanism, 530 potential, 534 UreE, 556 Metalloenzymes (see also Enzymes and individual names) evolution, 373, 392, 393 Metallothioneins, 47, 588, 631 Meteorites nickel in, 4 Methane 14 C, 346 acidity constant, see Acidity constants anaerobic oxidation, 343–346, 349, 350, 525 formation, see Methanogenesis Methanobacterium thermoautotrophicum (see also Methanothermobacter marburgensis), 324 Methanocaldococcus jannaschii, 370, 371, 373, 382, 384, 387, 388 Methanococcoides burtonii, 389

SUBJECT INDEX

Methanococcus jannaschii, 509 maripaludis, 327, 371, 387 thermolithotrophicus, 509 vaniellii, 387 voltae, 338 Methanogenesis, 525, 526 aceticlastic, 384–389 acetyl-coenzyme A decarbonylase/ synthases, 360, 384 F430, 182, 185, 188, 190, 325, 326, 332–334, 346–349 inhibition, 388 methylotrophic, 384, 386–389 Methanopyrus kandleri, 327, 338, 369–371, 373, 387, 388 Methanoculleus thermophilus, 338 Methanosarcina sp., 327, 338, 371, 526 acetivorans, 369, 370, 372, 384, 388, 389 barkeri, 338 mazei, 388, 389 thermophila, 359, 376, 384, 385, 388 Methanosphaera stadtmanae, 327, 389 Methanospirillum hungatii, 386, 387 Methanothermobacter marburgensis (see also Methanobacterium thermoautotrophicum), see Methylcoenzyme M reductase thermoautotrophicus, 371, 386 Methionine (and residues) S-adenosyl-, 475, 476, 481, 526, 528 formation, 474 salvage pathway, 229, 474–477, 496, 528 9-Methyladenine, 138 -1,3-dideaza-, 144 Methylation of cytosine, see Cytosine DNA, see DNA histones, 584 3-Methylcholanthrene, 585 Methylcobalamin, 211, 476 Methyl-coenzyme M, 182–185, 189, 190, 331, 334, 339, 347, 348 models, 334, 335

SUBJECT INDEX

Methyl-coenzyme M reductase (see also F430), 34, 37, 182, 323–350, 453, 525, 526, 529 active site, see Active sites catalytic cycle, 332, 333, 347–349 catalytic properties, 345–350 dual-stroke engine mechanism, 349, 350 inhibitors, 331, 345, 349 γ -irradiation, 342 isoenzymes, 324, 327, 328, 335, 346 mechanism, 183–185, 190 molecular properties, 335–345 Ni(I), see (Ni) Ni(III), see Ni(III) reactions, 335 spectroscopic studies, 337–341 structures, 334, 335, 338 sulfur coordination, 341, 342 Methylglyoxal as inhibitor, 449 cellular effects, 448, 449 cytotoxicity, see Cytotoxicity degradation, 449, 450 detoxicification, 449, 450, 465 formation, 446–448, 464 metabolism, see Metabolism synthase, see Synthases Methyl group transfer, 204, 360, 475, 526 acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 377, 379, 382 corrinoid iron-sulfur proteins, 383, 392 to Ni(I), 333 Methylmalonyl-CoA epimerase, 463 Methyltetrahydrofolate, 360, 382, 383, 390 Methylthioadenosine, 475, 483, 528 formation, 476, 481 homeostasis, see Homeostasis metabolism, see Metabolism nucleosidase, 477 phosphorylase, 477 Methylthioethyl sulfonate, see Methylcoenzyme M Methyl thiophosphate acidity constants, 155 complexes, 154–156 stability constants, 155, 156

685

Methyltransferases, 382, 475 5-Methyluracil, see Thymine Methyl viologen electron transfer, 203 Metronidazole, 549, 568 Mexico nickel concentrations, 14 Mice acireductone dioxygenase, 475, 483–486, 494 glyoxalases, 451 nickel studies, 585–588, 591, 602, 604, 607, 625, 628–631, 639–642 sperm, 84 Microarray analysis of DNA, 389, 584, 587–589, 605, 607, 608 GeneChip, 596, 597 Micrococcus ureae, 273 Microconstants, see Acidity constants and Stability constants Micronutrients for cyanobacteria, 37–40 plants, 37–40 Microorganisms (see also individual names and species) anaerobic, 326 pathogenic, 521, 527 Minerals (see also Ores and individual names) silicate, see Silicate Mining of nickel, 4, 6, 7, 41, 583 Mitochondria synthesis for [FeS] clusters, 287 Mitomycin resistance protein, 463 Mixed ligand complexes (see also Ternary complexes), 132 stack formation, 164, 165 with nucleotides, 159–165 Models for acetyl-coenzyme A synthase, 203–211 active sites of nickel enzymes, 181–230 carbon monoxide dehydrogenases, 200–211 cofactor F430, see F430 [NiFe] hydrogenases, 191–199

686

Molecular chaperones, 520, 529, 531, 534–537 acetyl-coenzyme A synthase, see Acetyl-coenzyme A synthases assist in re-folding, 534 ATP-dependent, 524 carbon monoxide dehydrogenases, see Carbon monoxide dehydrogenases GTP-dependent, 521, 523, 524, 533, 536, 538 hydrogenases, see Hydrogenases mechanism, 535 prevent misfolding, 534 SlyD, see SlyD urease, 535–537 Molecular mechanics calculations, 187 F430, 329 Molecular switch, 271 Molybdate, 267 Monkeys nickel studies, 628 Monoamine oxidases, 448 Moorella thermoacetica, 200, 359, 369, 371, 373, 375, 376, 390, 525 Moss as biomonitor for nickel, 8, 9, 11, 17 metal uptake efficiency, 8, 9 Mössbauer spectroscopy acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 376, 377, 379 C-cluster of carbon monoxide dehydrogenases, 364, 395 [NiFe] hydrogenases, 192, 282 Mus musculus (see also Mice), 474 Mutagenesis carbon monoxide dehydrogenases, 363 glyoxalase I, 462 nickel compounds, 584–587, 643, 644 nickel superoxide dismutase, 425, 426 [NiFe] hydrogenases, 287, 289–291, 298, 532, 533 site-directed, 212, 291, 298, 381, 530, 531 urease, 536 Mycobacterium tuberculosis, 529

SUBJECT INDEX

N NAD⫹, 280, 283 NADH ubiquinone oxidoreductase, 283 NADP, 280 NADPH, 449 Nanotechnology, 168 Neisseria meningitidis, 451, 453, 460–462 Nervous system nickel toxicity, 629 Neurodegenerative diseases (see also individual names), 418 Niccolite, 6 Nickel (different oxidation states) (in) 60 Ni, 3 61 Ni, 34, 195, 206, 281, 301, 303, 342, 429, 431 63 Ni, 39, 626, 639 allergy, see Allergy and Helicobacter pylori, 545–570 antimonides, see Antimonides arsenides, see Arsenides atmosphere, see Atmosphere binding by strong ligands, 46, 47 bioavailability, see Bioavailability biogeochemistry, see Biogeochemistry biological function of hyperaccumulation, 49, 50 –carbon bonds, see Bonds carcinogenesis, see Carcinogenesis carriers, see Transporters catalyst of early life, 35–37 chemistry, 2–6, 33–35, 64, 111 chloride, see Nickel chloride citrate, see Citrate -containing jewelry, 624 coordination numbers, 34, 35 coordination sphere, 196, 197, 251, 252, 421 cyanobacteria, see Cyanobacteria deficiency, see Deficiency -dependent gene expression, 581–609 environment, see Environment exclusion, 45, 46 excretion, 621, 622, 625–627

SUBJECT INDEX

[Nickel (different oxidation states) (in)] exposure, 84, 582–585, 602, 621–624, 638, 645 fluxes in forests, 12–14 geometry, 34, 35, 65, 114, 166, 182, 474 health effects of exposure, 582–585, 597 historical records of deposition, 14–16 homeostasis, see Homeostasis hydrogenases, see [NiFe] hydrogenases hydroxide bridge, see Hydroxide hyperaccumulators, see Hyperaccumulation in plants incorporation into proteins, 528–530 -induced changes in calcium homeostasis, 602–604 -induced expression of erythropoietin, 592, 593 interdependency with other metal ions, see Interdependencies isotopes, 3, 34 lakes, see Lake mechanisms of regulation, 562–566 metallurgy, 621 ores, 4 oxidation states, 34, 35 oxides, see Oxides pathologic effects, 623 phosphate bridge, 262, 263 plants, see Plants pollution, 40–48 Raney, 34 redox processes, 3 refinery workers, 623, 638 resistance in plants, 45–48 responsive regulator NikR, 552–554 sample measurements, 4–6 seawater, 110 sequestration, 45, 46 serum albumin, 84 silicates, see Silicate soil, see Soil sulfates, see Sulfates sulfide, see Sulfides sulfur-rich sites, 191–214 tetrahydrocorphinoid cofactor, see F430 toxicity, see Toxicity

687

[Nickel (different oxidation states) (in)] transporters, see Transporters uptake, 560, 561, 621–627 uses, 6 water, see Lake and Water Nickel(0) (in), 200, 204, 209, 330 acetyl-coenzyme A synthase, 377–379 Ni(II)Ni(0), 210 Nickel(I) (in), 34, 183 acetyl-coenzyme A synthase, 204, 205, 377–379 conformational change, 340 coordination sphere, 187 ESR studies, see EPR F430, 184, 185, 188, 328–333, 337–341, 345, 347–349, 525 hydroporphyrins, 330 methyl-coenzyme M reductase, 335, 339 [NiFe] hydrogenases, 302, 311 Ni(II)Ni(I), 210 porphyrin-type ligands (see also Porphyrins), 186 redox potential, see Redox potentials reduction, 330 Nickel(II) (in/complexes with) amino acids, 63–97 aminohydroxamate, see Aminohydroxamate aminophosphonate, see Aminophosphonate chemical properties, 111 coordination sphere, 114, 166, 182, 474 disulfide, see Disulfide -EDTA, see EthylenediamineN,N,N⬘,N⬘-tetraacetate hormones, see Hormones mechanisms of toxicity, 600 multicomponent systems, 94–96 Ni(II)Ni(0), 210 Ni(II)Ni(I), 210 nucleic acid binding, 165–168 nucleobase residues, 118–128 nucleotides, 109, 148–150, 169 peptides, 63–97 phosphate, 128–130

688

[Nickel(II) (in/complexes with)] purine-nucleotide (N1)-oxides, 150–154 purines, 118–122, 146 pyridine derivatives, 126 pyrimidines, 122–128 redox potential, see Redox potentials stability constants of complexes, see Stability constants sugars, 112–117 ternary ATP–buffer complexes, 161–163 ternary nucleotide complexes, 164 thioethers, see Thioether thiophosphate, 155, 156 uridine, 127, 128 water exchange rate, 111 Nickel(III) (in/complexes with), 34, 183 -alkyl bond, see Bonds allergenic, 629 bleomycin, 92 compounds, 190 coordination sphere, 39 cyclam, 144 EPR, see EPR F430, 188, 329, 331–333, 341, 342, 349 methyl-coenzyme M reductase, 340–342 [NiFe] hydrogenases, 300–303, 307–312 peptides, 64, 86–94, 209 photoreduction, 426–428 reduction, 212, 434 stabilization, 65, 66 superoxide dismutase, see Nickel superoxide dismutase ternary complexes with peptides, 92 thiolate, 341 Nickel(IV), 90 allergenic, 629 Nickel carbonyl, 646 14 C-labeled, 626 63 Ni-labeled, 626 detoxification, 624 inhalation, see Inhalation toxicity, see Toxicity

SUBJECT INDEX

Nickel chaperones, 248, 286, 367, 519–539 carbon monoxide dehydrogenase, see Carbon monoxide dehydrogenase mechanism, 530 metallo-, see Metallochaperones molecular, see Molecular chaperones [NiFe] hydrogenases, see [NiFe] hydrogenase SlyD, see SlyD urease, see Urease Nickel chloride, 628–630, 635, 637 injection of 63Ni chloride, 626 oral exposure to 63Ni chloride, 626 Nickel-containing enzymes, see individual names Nickel iron hydrogenases, see [NiFe] hydrogenases Nickel subsulfide, 585, 592, 593, 602–604, 606, 607, 629, 630, 633, 635, 636 35 S-labeled, 627, 639 63 Ni-labeled, 627, 639 carcinogenic potential, 646 inhalation, see Inhalation Nickel superoxide dismutase, 39, 182, 414–438, 526 activation, 527 active site, see Active sites crystal structures, 428, 429, 526 enzymes, 418–422 inhibitors, 432 kinetics, 434–436 mechanism, 429–437 models, 211–214, 436, 437 molecular biology, 422–426 mutants, 423, 431 Ni(III), 422, 426, 429, 434, 436, 526 redox potential, see Redox potentials reduction, 426–428, 433 sequence alignments, 424 spectral parameters, 432 spectroscopy, 429–434 Streptomyces sp., 422–438 structural biology, 426–429 wild-type, 423, 425, 427, 431, 432

SUBJECT INDEX

Nicotianamine, 38, 47 Nicotiana tabacum, 47 Nicotinamide adenine dinucleotide, see NAD⫹ Nicotinamide adenine dinucleotide (reduced), see NADH Nicotinamide adenine dinucleotide phosphate, see NADP Nicotinamide adenine dinucleotide phosphate (reduced), see NADPH [NiFe] hydrogenases, 35–38, 81, 97, 182, 279–314, 421, 514, 521, 523 A. vinosum, 295, 297–299, 305, 307–309, 311 active site, see Active sites bacterial, 283 biosynthesis, see Biosynthesis carbon monoxide in, see Carbon monoxide catalytic cycle, 281, 282, 295, 296, 298, 307, 309–313 coordination geometry, 291, 301 (crystal) structures, 283, 284, 291–294 cyanide, see Cyanide Desulfovibrio sp., 196, 199, 281, 283, 285, 287, 289, 290–302, 304–306, 312, 421 electrochemistry, see Electrochemistry electron transfer, see Electron transfer electron transport mutations, 289 energy-converting isoenzyme I, 326 Escherichia coli, see Escherichia coli eukaryotic, 287 F420 -nonreducing, 325 F420 -reducing, 325 Fe(II), 193, 309–311 gas channel, 281, 282, 289, 290, 293–295, 312, 313 genes, 284–291 genetic manipulation, 287–291 Helicobacter pylori, 559, 560 inhibition, 294, 308 isotope-labeled, 292, 301–305 maturation pathway, 285–288 mechanism, 192 metallochaperones, 532, 533

689

[[NiFe] hydrogenases] models, 191–199, 212 Ni(I), 302, 311 Ni(III), 300–303, 307–312 oxygen tolerance, 290 prokaryotic, 284, 285, 287 proton transfer, see Proton transfer proton transport mutations, 289, 290 Ralstonia eutropha, 285–287, 289–291, 295, 300, 304, 306, 307, 309 redox states, 296, 297 schematic view, 282 single crystal, 293, 301, 302, 305 spectroscopic measurements, 192, 193 spectroscopic studies, 295–306 Thiocapsa roseopersicina, 285, 287 wild-type, 290, 291 Nigeria nickel concentrations, 10 Nitellopsis, 41 Nitric oxide, 551 Nitric oxide synthase inducible, 551, 590, 597 Nitrilotriacetate Ni(II) complex, 81, 118, 167 Nitrogen 14 N, 306, 431, 432 15 N, 301, 306, 431, 432 deficiency, see Deficiency fixation, 38 metabolism, see Metabolism Nitrogen monoxide, see Nitric oxide 4-Nitrophenylphosphate hydrolytic cleavage, 217 NMR (studies of) 1 H, 86, 151, 198, 480, 488 13 C, 488, 489 15 N, 488, 489 acetyl-coenzyme A synthase, 209 acireductone dioxygenase, 474, 480, 488, 490 F430, 333 hormone complexes, 86 9-[2-(phosphonomethoxy)ethyl]adenine, 151 ternary nucleotide complexes, 164

690

Norway Arctic, see Arctic nickel concentrations, 8, 9, 12–14 NTA, see Nitrilotriacetate Nuclear magnetic resonance, see NMR Nucleic acids, see DNA and RNA Nucleobases (see also individual names) acidity constants, see Acidity constants Ni(II) complexes, 118–128 oxidation, 111 stability constants, see Stability constants structures, 118 Nucleophilic attack (of), 491 Co(I), 383 cysteine, 296, 313 hydroxide, 200, 215–218, 269–271 Ni(I)-F430, 184 Nucleoside(s) (see also individual names) phosphates, see Nucleotides and individual names self-association, see Self-association 5⬘-O-thiomonophosphate complexes, 154–156 triphosphates, see individual names Nucleoside diphosphate(s) (see also individual names) acidity constants, see Acidity constants complexes, 139–144 purine-, 139 pyrimidine, 139 Nucleoside monophosphate(s) (see also individual names) 2⬘-, 144–146 3⬘-, 144–146 acidity constants, see Acidity constants Ni(II) complexes, 113–115, 132–139, 144–146 purine-, 135–139 pyrimidine-, 134, 135, 138 Nucleoside triphosphate(s) (see also individual names) complexes, 139–144 dephosphorylation, 144 Nucleotide(s) (see also individual names) (complexes with) acidity constants, 115, 156 acyclic analogs, 156–158

SUBJECT INDEX

[Nucleotide(s)] derivatives, 149–159 less common ones, 144–149 Ni(II), 109–169 oligo-, 165 with a Pt(II)-coordinated nucleobase, 158, 159 Nutrients micro-, see Micronutrients

O Oil combustion, 7 nickel in, 7, 21 OMP, see Orotidine 5⬘-monophosphate Ores nickel, 4 Origin of life, 392, 393 Ornithine decarboxylase, 476 Ni(II) complex, 69, 71 Orotidine 5⬘-monophosphate acidity constant, 146 Ni(II) complex, 146 structure, 147 Oryza sativa, 481, 483, 486 Osteosarcoma human, 606 Ovarian cancer, see Cancer Oxalate decarboxylase, 493, 494 oxidase, 493, 494 Oxidases (see also individual names) copper amine, 493 monoamine, see Monoamine oxidase oxalate, 493, 494 Oxidative damage (by/in) DNA, 600, 643, 644, 646 effect of nickel, 633–637 methylglyoxal, 448, 449 proteins, 600 superoxide, 418, 526 Oxidative stress, 390, 587, 589, 601, 605 in plants, 42, 44, 45, 47, 48, 51

SUBJECT INDEX

691

[Oxidative stress] nickel-induced, 599–602, 644, 646 resistance, 554 Oxides aluminum, 13 iron, 13, 20 manganese, 20 nickel, 8, 636, 639, 642 Oxidoreductases (see also Peroxidases and individual names) cytochrome c3, see [NiFe] hydrogenase NADH:ubiquinone, 283, 326 superoxide dismutases, see Superoxide dismutases Oximes α-alanine, 75 derivatives of amino acids, 75, 76 Ni(II) complexes, 71, 75, 76 6-Oxopurine, see Hypoxanthine Oxygen (see also Dioxygen) 17 O-labeling, 292, 301, 305 18 O, 229 production, 308, 314 Oxytocin Ni(II) complex, 78

P Palladium(II) purine-nucleobase complexes, 121, 145 Parkinson’s disease, 86, 418 Parvulins, 503–505 Pathogens (see also Bacteria and individual names) fungal, see Fungi gastric (see also Helicobacter pylori), 545–570 plant, see Plants Peas nickel in, 19 Peat (containing) lead, 15 mercury, 15 nickel, 5, 7, 10, 11, 14–16 pre-anthropogenic, 16 scandium, 16

PELDOR, see Pulsed electron-electron double resonance Pelobacter carbinolicus, 391 Peptide(s) (see also Amides and individual names) (containing/ complexes with) amide functions, 76 bond hydrolysis, 83 Cd(II), 80 Cu(II), 76, 77, 79, 80 Cu(III), 87, 93 cyclo-, 81 fibrino-, 84 fragments from histones, 82–86 fragments from protamines, 82–86 hormones, 78, 85 involved in catalytic ractions, 86–90 involved in redox reactions, 86–90 Ni(II), 63–97 Ni(III), 86–94, 209 octa-, 84 sulfhydryl, 81 thiolate, 81, 82 with coordinating side chains, 77–80 with cysteinyl residues, 80–82 with histidyl residues, 80–82 with non-coordinating side chains, 76, 77 Zn(II), 77, 80 Peptidyl prolyl cis/trans isomerases (see also individual names) activity, 533 cyclophilins, see Cyclophilins Escherichia coli, see Escherichia coli FK506-binding protein, see Proteins Ni(II)-mediated inhibition, 515 Ni(II)-regulated, 501–515 parvulins, 503–505 SlyD, see SlyD Permeases nickel, 39, 286 Permittivity, see Dielectric constant D -Penicillamine nickel detoxification, 624 Peridotite, 4 Peroxidases ascorbate, 44 glutathione, 628, 636, 637

692

Peroxide(s), 308, 418, 526 lipid, 646 hydro-, see Hydroperoxides hydrogen, see Hydrogen peroxide Peroxynitrite, 551 Persulfide, 365 bond, 370 formation, 371 Phaseolus vulgaris, 44 1,10-Phenanthroline (and derivatives), 376 Ni(III) complex, 92 ternary complexes, 164 Phenolate bridge, 219 Phenylalanine Ni(II) complex, 68, 71 Phenylglyoxal, 446, 451 Phenylphosphorodiamidate, 256 hydrolysis, 268, 270 Phosphate(s) (see also individual names) acidity constants, see Acidity constants as urease inhibitor, 261–263, 267–272 di-, 128–130 mono-, 128–130 Ni(II) complexes, 128–130 stability constants, see Stability constants tri-, 128–130 Phosphines, 196, 202, 204, 208, 210 Phosphinic acid, 79 Phosphohydrolase ATP, 369 Phospholipids, 589 synthesis, see Biosynthesis Phosphonate(s) (or phosphonic acid), 79, 134, 136 acetonyl-, see Acetonylphosphate amino-, see Aminophosphonates 9-[2-(Phosphonoethoxy)ethyl]adenine complexes, 158 9-[2-(Phosphonomethoxy)ethyl]adenine complexes, 136, 156–159 diphosphorylated, 156 structure, 151 ternary complexes, 164 Phosphorus 32 P, 512 Photosynthesis inhibition, 43–45, 51

SUBJECT INDEX

Photosystem I inhibition, 43 Photosystem II inhibition, 43, 44 Phyllanthus pallidus, 53 Phylogenetic tree acetyl-coenzyme A decarbonylase/ synthase, 371, 372, 381, 382 acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 371–373, 381, 382 carbon monoxide dehydrogenases, 373 methyl-coenzyme M reductase, 327 Phytochelatin, 47, 51 Phytomining, 33, 48, 49, 52, 53 Phytoplankton, 32, 39 Phytoremediation (see also Hyperaccumulation in plants), 33, 48, 52, 53 Picea glehnii, 46 Picolinamide cleavage, 218 stability constants, see Stability constants Pigments nickel in, 6 Pine copper uptake, 20 nickel-excluding, 46 nickel uptake, 12, 17, 19, 20, 40, 45 Ping-pong mechanism in acetyl-coenzyme A decarbonylase/ synthase, 385 nickel superoxide dismutase, 418, 434, 435, 437, 438 Pinus sylvestris, 10, 18, 40, 46 Pisum sativum, 44 Plant(s) (see also individual names and species) copper uptake, 41, 43 dose-response relationship for metals, 32 excluders, see Excluders geneticall engineered, 53, 54 glyoxalases, 451 growth inhibition, 33, 42–45 higher, 39, 41–44

SUBJECT INDEX

[Plant(s)] hyperaccumulators, see Hyperaccumulating plants inhibition of photosynthesis, 43, 44, 51 inhibition of root function, 42–44 iron superoxide dismutase, 418 iron uptake, 43 metabolism, see Metabolism micronutrients, see Micronutrients nickel concentrations, 12, 17–19 oxidative stress, see Oxidative stress pathogens, 40, 49 resistance against nickel toxicity, 45–48 trace elements in, 32, 43 urease, 242, 554 Plasmodium sp., 451 falciparum, 460 Platinum(II) (complexes with) -coordinated nucleobases, 158, 159 diethylenetriamine, 122 purine-nucleobases, 121, 145 pyrimidine, 122 Pleurozium schreberi, 9 PMEA, see 9-[2-(Phosphonomethoxy)ethyl]adenine Polar ice, 15, 16 snow, 15, 16 Pollutant degradation, 524 Porphyrins (see also individual names) Ni(I), 183, 186, 188 Ni(II), 187 Prion proteins octapeptide repeats, 84 Prochlorococcus marinus, 39, 527 Prokaryotes (see also individual names), 477, 481, 526 FK506-binding proteins, 503, 509 glyoxalases, 450, 465 iron superoxide dismutase, 418 manganese superoxide dismutase, 418, 419 [NiFe] hydrogenases, see [NiFe] hydrogenases SlyD, 505

693

Proline (and residues), 85 hydroxylation, 593–598, 646 Ni(II) complex, 68 Prolyl bond cis/trans isomerization, 501–515 Propanoate 2-hydroxyimino-, 75, 76 Prostate tumor, see Tumor Prostheses nickel allergy, 592, 622, 638 Protamines (binding to) compact DNA binding, 84 Cu(II), 84 Ni(II), 82, 84–86, 90, 632, 634 Protein(s) (see also Enzymes and individual names) AP-1, 644 Ca(II) binding, 85 cleavage, 90 damage by nickel, 631, 634, 635 DNA recognition, 90 extracellular matrix, 590, 591 FK506-binding, 502–505, 509–511, 513, 515, 533 folding, 503 fosfomycin resistance, 463, 464 Fur, 553, 564–566 HIF-1, 644 high-potential iron, 422 hydrogen-sensing, 284, 287, 290, 291 hydrophobic environment, 83 hydroxylation, 593–599 –ligand interactions, 503 NF-κ B, 644 nickel incorporation, 528, 529 nickeloplasmin, 631 NikR, 552–554, 562–566 NixA, 566 –protein interaction, 284, 503, 593, 595 RNA recognition, 90 surfactant, 588–590 synthesis, see Synthesis transporters, see Transporters UreE, 556 Protein Data Bank (files of protein structures) acetyl-coenzyme A synthase, 523

694

[Protein Data Bank (files of protein structures)] acireductone dioxygenases, 475, 481, 483–485, 494, 523 carbon monoxide dehydrogenases, 36, 523 glyoxalase, 523 methyl-coenzyme M reductase, 523 nickel superoxide dismutase, 523 [NiFe] hydrogenases, 36, 523 peptidyl prolyl cis/trans isomerases, 509 ureases, 246, 247, 249–253, 255–268, 523 Proteus mirabilis, 255, 530, 555 Protonation constants, see Acidity constants Proton transfer carbon monoxide dehydrogenase, 362, 363, 370, 371 -coupled electron transfer, see Electron transfer mutations in hydrogenases, 289, 290 [NiFe] hydrogenases, 282, 293, 298, 312, 313 Pseudomonas aeruginosa, 451, 453, 460–462, 464, 553 putida, 451, 452, 460–462, 527 Pterins sarcina-, 382 tetrahydromethano-, see Tetrahydromethanopterin Pulsed electron-electron double resonance [NiFe] hydrogenase, 301 Pulse radiolysis nickel porphyrins, 188, 190 Purine(s) (see also individual names) 6-hydroxy-, see Hypoxanthine N9-substituted, 118 Ni(II) complex, 118–122 nucleotides, see individual names self-association, 132 stacking, see Stacking Putrescine, 476 Pyridine(s) acidity constant, 120 as ligand, 123, 124, 126, 127 o-amino-, 123, 124, 126, 127

SUBJECT INDEX

Pyrimidine(s) (see also individual names) (complexes with) Ni(II), 122–128 nucleoside 5⬘-monophosphates, 134, 135 nucleoside 5⬘-triphosphates, 141, 143 Pyrococcus horikoshii, 509, 529 Pyruvate, 12, 446, 447

Q Quercetin 2,3-dioxygenase, 493–495 Cu(II), 493, 494 Fe(II), 493–495 structure, 494

R R. albus, see Ruminococcus eutropha, see Ralstonia leguminosarum, see Rhizobium palustris, see Rhodopseudomonas rhodochrous, see Rhodococcus rubrum, see Rhodospirillum Rabbit nickel studies, 628–631, 641 Radicals (see also individual names), 479, 489, 495 alkyl, 188, 190 allyl, 349 disulfide, 348 free, 334, 348 hydroxyl, 90, 551, 633 intermediates, 634 methyl, 334, 348, 349 Ni(II)-peptide intermediate, 89 sulfuranyl, 185, 333 superoxide, see Superoxide thiyl, 185, 333, 341, 342, 348, 349 Radiolysis pulsed, see Pulsed radiolysis Ralstonia eutropha, 285–287, 289–291, 295, 300, 304, 306, 529

SUBJECT INDEX

Raman spectroscopy (studies of) DNA, 167 methyl-coenzyme M reductase, 341 nickel superoxide dismutase [NiFe] hydrogenase models, 199 Ranithidine bismuth citrate, 549 Rat acireductone dioxygenase, 481, 483 glyoxalase I, 451 nickel studies, 585, 592, 593, 604, 606, 625, 626, 628–631, 639–642 Rate constants superoxide dismutases, 420, 434 urea hydrolysis, 226 water exchange, 111 Reactive oxygen species, 111, 418, 551, 552, 583, 588, 600, 602, 633–637, 643, 644, 646 detoxification, see Detoxification Redox potential (of) carbon monoxide dehydrogenase, 361, 364 CO/CO2, 387 disulfide, 346 F430, 330–333 [Fe4S4] cluster, 361 Ni(II)/Ni(I), 183, 186, 190, 330, 334, 348, 349, 419 Ni(III)/Ni(II), 88, 91, 92, 94, 193, 212, 331–333, 348, 419, 421, 422, 434, 438, 633 Ni2⫹, 490 [NiFe] hydrogenases, 198, 297, 298 [Ni(His)]⫹, 71 thiyl radical/thiol couple, 348 Reductases glutathione, 628 superoxide, 422 Reduction potential, see Redox potential Resistance antibiotics, 568 fosfomycin, 463, 464 nickel, 45–48 oxidative stress, 554 Respiratory tract cancer, see Cancer nickel toxicity, 628

695

Retinoblastoma, 606 Rhamnose Ni(II) complex, 113 Rhizobium sp., 19 leguminosarum, 533 Rhodium(I) 8-azaadenine complex, 146 Rhodococcus rhodochrous, 529, 560, 561 Rhodopseudomonas palustris, 368, 391 Rhodospirillum rubrum, 200, 359, 387, 390, 391, 524, 534, 539 Ribose 2⬘-deoxy, 112 5-monophosphate, 130 acidity constant, 112 hydroxyl groups, 112, 115 methylthio-, 477, 479 Ribozymes, 122 probes, 154 River (see also Water) nickel-polluted, 41 RNA cleavage, 167 Ni(II) complex, 167 recognition, 90 synthesis, see Synthesis tobacco mosaic virus, 448 Ruminococcus albus, 390 Russia Kola Peninsula, see Kola Peninsula nickel mining and smelting, 4, 8, 9–11, 14, 18, 20, 42

S S. aureus, see Staphylococcus cerevisiae, see Saccharomyces coelicolor, see Streptomyces fumoroxidans, see Syntrophobacter lividans, see Streptomyces pombe, see Schizosaccharomyces salivarius, see Streptococcus seoulensis, see Streptomyces thermophilus, see Streptococcus

696

[S.] typhimurium, see Salmonella xylosus, see Staphylococcus Saccharomyces cerevisiae (see also Yeast) acireductone dioxygenase, 481 glyoxalases, 448, 449, 451, 460–462, 527 Salen (complexes with) Ni(II), 167 Ni(III), 89 Salmonella sp., 585 typhimurium, 643 Sample limit of detection, 4, 5 nickel, 4–6 Sarcomas (see also Tumors) nickel-induced, 641 Sarcosine hydroxamic acid, see Hydroxamic acid Scandinavia, 20 Scandium in peat, see Peat Scenedesmus sp., 47 acutus, 41, 45 obliquus, 291 Schiff bases (see also individual names) (complexes with) Ni(II), 71 Ni(III), 92 Schizosaccharomyces pombe, 529 SDS-PAGE glyoxalase I, 453 Seawater nickel in, 5, 110 Sebertia acuminata, 47, 48, 51 Secreted protein, acidic and rich in cysteine, 85 Sediments anoxic, 3 nickel in, 14, 15 Selenium 77 Se, 301 Selenocysteine (and residues), 280, 292, 523 Self-association (see also Stacking) nucleosides, 139 Semichelates, 123, 126, 128, 130 Senecio coronatus, 49

SUBJECT INDEX

Serine α-hydroxymethyl-, 79 Ni(II) complex, 69 Serpentine, 4, 33, 48 Sewage sludge, 2, 21 Siderophores (see also individual names), 72, 553 Signaling pathways, 584 alteration, 593–607 Signal transduction, 144, 280, 644 Silene paradoxa, 41, 43 Silicate minerals, 15 nickel, 51 Skin dermatitis, see Dermatitis inflammation, 629 nickel exposure, 622, 624 SlyD (see also Peptidyl prolyl cis/trans isomerases) affinity for Ni(II), 512 biological role, 513, 514 catalyzed reactions, 503, 504 chaperone-like activity, 511, 532, 533 distribution of genes, 505–508 Escherichia coli, see Escherichia coli inhibition by nickel, 511, 512 inhibition of adenylate cyclase, 513 link to hydrogenase biosynthesis, 514 Ni(II)-regulated, 501–515 nucleotide binding, 512, 513 sequence alignment, 510 structure, 508–510 Zn(II), 512 Smelting, 41 copper, 8–11, 15, 17, 18, 20, 42 nickel, 7–11, 14, 15, 17, 18, 20, 21, 41, 42, 583 Sodium ion channel, 629 diethyldithiocarbamate, 624 Sodium dodecyl sulfate electrophoresis, see SDS-PAGE Soil (containing) contaminated, 48, 52, 53 nickel mobility, 17–21

SUBJECT INDEX

[Soil (containing)] nickel, 2–4, 12, 33, 42, 45 urease, 242 Solvent polarity reduced, 115–117 Sorbose Ni(II) complex, 113 Soybean (see also Glycine max.), 8, 451 nickel in, 18, 19 SPARC, see Secreted protein, acidic and rich in cysteine Sperm abnormalities, 630 human, 84 mouse, 84, 630 vertebrate, 84 Spermidine N⬘-monoglutathionyl-, 464, 465 synthesis, 476 Spermine synthesis, 476 Spirodela, 41 Sporosarcina ureae, 555 Spruce nickel concentration, 12 Stability constants of complexes (see also Equilibrium constants) apparent, 152–154 5⬘-ATP, 143, 162, 163 buffers, 163 conditional, 70 Cu(II)-nucleotides, 140, 142, 150 GMP, 137 methyl thiophosphate, 155, 156 Mg(II)-nucleotides, 136, 142, 143 microconstants, 154 Mn(II)-nucleotides, 142, 143, 150 Ni(II)-nucleotides, 116, 117, 134–143, 148, 162, 163 Ni(II)-phosphate monoesters, 116, 117, 136 Ni(II)-purine-nucleobases, 119, 120 Ni(II)-pyrimidines, 123–125 Ni(III)-peptides, 92 nickel-amino acids, 66–68 nickel-picolinamide, 218 purine-nucleotide (N1)-oxides, 152–154

697

[Stability constants of complexes] phosphates, 129, 130 ternary ATP-buffer complexes, 162, 163 Zn(II)-nucleotides, 142, 143 Stackhousia tryonii, 50 Stacking (see also Self-association) aromatic ring, 165 intramolecular, 141, 164, 165 Ni(II)-peptide complexes, 78 purines, 132, 139 pyrimidines, 139 Stainless steel nickel in, 21 production, 6 Standard potential, see Redox potential Staphylococcus aureus, 448 xylosus, 266 Stereocaulon paschale, 10 Stomach, 546 Stopped flow study of urease, 220 Streptanthus insignis, 49 polygaloides, 49, 52 Streptococcus salivarius, 556 thermophilus, 556 Streptomyces sp., 39, 211, 422–426, 428, 436, 527 coelicolor, 214, 423–425, 428–430, 526 lividans, 423 seoulensis, 423–425, 428, 429, 431, 526, 527, 534 Strontium(II) flavin mononucleotide complex, 148 xanthosine 5⬘-monophosphate complex, 148 Sugars, 113 Ni(II) complexes, 112–117 Sulfate, 267 and urease, 267 nickel, 8, 588–590, 606, 623, 628–630, 636, 638, 640, 643 reduction, 389, 391 Sulfhydryl groups, 81 Ni(III) complexes, 92

698

Sulfide(s) bridge, 36, 366, 367, 524 ligand, 422, 426 nickel sub-, see Nickel subsulfide nickel, 3, 4, 8, 15, 35, 586, 600, 633, 638 ores, 4 per-, see Persulfide poly-, 340–342 Sulfoxide in [NiFe] hydrogenases, 294 Sulfur (different oxidation states) 33 S, 340, 342, 431, 639 in melanin, 296, 297 oxidation, 633 Sulfur dioxide, 9, 10 Superoxide, 211, 212, 214, 418–420, 526, 551, 646 formation, 490, 491 oxidation, 437 reductase, see Reductases reduction, 428 Superoxide dismutase(s), 551, 553, 554, 600, 637 Cu/Zn, see Copper-zinc superoxide dismutase iron, see Iron superoxide dismutase manganese, see Manganese superoxide dismutase nickel, see Nickel superoxide dismutase redox potential, see Redox potential Sweden nickel mining, 6 Synechocystis sp., 39 Synthases carbamoyl phosphate, 286 glutathione, 600 methylglyoxal, 446–448 Synthesis (see also Biosynthesis) (of) DNA, 449 protein, 449 RNA, 449 spermidine, 476 spermine, 476 urea, 273 Syntrophobacter fumaroxidans, 370, 382

SUBJECT INDEX

T Taiwan, 15 Tannins, 45, 46 Ternary complexes (of) (see also Mixed ligand complexes) Ni(II), 161–164 Ni(III), 92 1,4,8,11-Tetraazacyclotetradecane, see Cyclam Tetracycline, 568 Tetraglycine Ni(III) complex, 90 Tetrahydrofolate methyl-, see Methyltetrahydrofolate Tetrahydromethanopterin, 383 Tetrahydrosarcinapterin, 360 Tetrapyrroles, 525, 526 F340, see F430 Thermus thermophilus, 502 Thiocapsa roseopersicina, 285, 287 Thiocyanate (in) carbon monoxide dehydrogenases, 366 N-bound, 218 Thioether bond cleavage, 190 Ni(II) complex, 78, 79 7-Thioheptanoylthreoninephosphate (see also Coenzyme B), 325, 337–340, 345–349, 525 Thiols (and thiolate groups) (see also individual names) Ni(II) complexes, 78, 81, 83, 193, 194 Ni(III) coordination, 341, 342 nickel superoxide dismutase, 422, 426, 436, 437 Thiophosphate methyl-, see Methylthiophosphate Thiourea, 219 Thlaspi sp., 51, 52 caerulescens, 48, 50 calaminare, 48 goesingense, 38 Threonine Ni(II) complex, 69 Thrombin, 84

SUBJECT INDEX

Thymidine structure, 118 Thymidine 5⬘-diphosphate (complexes with) Cu(II), 140 Ni(II), 139, 140 Thymidine 5⬘-monophosphate (complex with), 133, 135 Ni(II), 134 Thymidine 5⬘-triphosphate metal ion complexes, 141, 143 Thymine 1-(2⬘-deoxy-β -D-ribofuranosyl)-, see Thymidine structure, 118 Thyrotropin releasing factor, 85 Titanium(III) citrate, 375, 377, 378 reduction of C-cluster of carbon monoxide dehydrogenase, 364 reduction of Ni(II)F430, 330, 331 Tobacco smoke, 638 transgenic plants, 47 Toxicity of copper in plants, 41, 42 cyanide, 623 Toxicity of nickel, 82, 587, 588, 600, 619–646 carbonyl, 622, 623 embryo-, 630, 631 geno-, 643–645 in experimental animals, 625–631 in plants, 17, 18, 33, 40–48, 51 mechanisms, 631–637 systemic, 627, 628 tissue, 628 Trace elements in plants, see Plants Transcription factors alteration, 593–607 AP-1, 598–603, 644 ATF-1, 599 HIF-1, 590, 592, 593, 595–598, 602, 603, 606, 644, 646 NF-κ B, 598–601, 644 zinc finger, 597

699

Transferases acetyl-, 608, 645 amino-, 600, 628 O-carbamoyl-, 286 glutathione-S-, 44, 600 Transferrin, 627 lacto-, 588, 589 Transition metal ions, see individual names Transporters (see also individual names) ABC-type, 528, 560 cobalt, 38, 39, 529 divalent metal, 595, 627 magnesium, 561 Ni/Co, 38, 39 nickel, 38, 39, 46, 560, 561, 566, 631 zinc, 38, 46 Triglycine Ni(II) complex, 76, 79 Triosephosphate isomerase, 447, 448, 460 Tris acidity constants, 163 stability constants of complexes, 163 structure, 162 ternary complexes, 161–163 Tris(hydroxymethyl)methylamine, see Tris Triticum sp., 43–45, 47 aestivum, 18 Trypanothione, 464, 465 Trypsin, 511 chymo-, see Chymotrypsin Trypsinogen, 602 Tryptophan (and residues) Ni(II) complex, 68 ternary complexes, 164 Tubercidin 5⬘-monophosphate, 144 Ni(II) complex, 134 Tumor(s) (see also Cancer, Sarcomas, and individual names) lung, 638, 639, 643 nickel-induced, 587, 588, 599 prostate, 483 suppressor gene, 606, 607 testicular, 640 Tumor necrosis factor, 592, 600, 601 Tyrosine Ni(II) complex, 68, 78

700

SUBJECT INDEX

U Ubiquinone NADH oxidoreductase, 283 5⬘-UDP Ni(II) complex, 139, 140 Ulcer duodenal, 546 gastric, 521, 548 Umbilicaria sp., 10 5⬘-UMP, 132, 133, 135, 146 UMPS, see Uridine 5⬘-Othiomonophosphate United Kingdom nickel in soil, 20 Uracil, 126 structure, 118 Urea, 219, 220, 272 acidity constants, see Acidity constants aminohydrolase, see Urease decomposition, 216, 243 half life, 243 hydrolysis, 226, 243, 249, 256, 265, 267, 268, 554, 556 production, 243 resonance stabilization energy, 223 synthesis, 273 thio-, see Thiourea Urease(s), 35, 37–40, 111, 241–273, 453, 514, 520, 521, 530 acetohydroxamic acid complex, see Acetohydroxamic acid acid resistance, 556–559 acidity constants, see Acidity constants activation, 521, 530–532, 537 active site, see Active sites apo-, 267 Bacillus pasteurii, 244–267 biochemistry, 243, 244 biological significance, 242, 243 carboxylate bridge, 249, 252 conformation, 248, 249, 261, 268, 272 decomposition, 554 electron density, see Electron density environmental adaptation, 554–556 gene cluster, 536, 555, 566

[Urease(s)] Helicobacter pylori, see Helicobacter pylori history, 37 hydroxamate bridge, 259 hydroxide bridge, see Hydroxide in Helicobacter pylori infection, 549, 552 inhibitors, 228, 255–263, 267–272, 569 jack bean, 242, 248, 251, 255, 261, 266, 267, 273 Klebsiella aerogenes, 244–267, 521, 536 mechanism, 215, 267–272 metallochaperones, 530–532 metal-substituted, 266, 267 models, 214–229 molecular chaperones, 535–537 mutants, 244, 259, 263–266, 271, 530, 531, 561, 567 Ni(II)-aminohydroxamate, 72 nickel insertion, 369 nickel regulation, 558, 562–566 phosphate bridge, 262 Proteus mirabilis, 255 Staphylococcus xylosus, 266 structure with a substrate analog, 255, 256 structure with a transition state analog, 256, 257 structures in the native state, 249–254 structures with a competitive inhibitor, 257–263 structures, 244–267 synthesis, see Biosynthesis water bridge, 252 wild-type, 265, 266, 567 Uridine complexes, 126, 127 structure, 118 Uridine 5⬘-diphosphate, see 5⬘-UDP Uridine 5⬘-monophosphate, see 5⬘-UMP Uridine 5⬘-O-thiomonophosphate acidity constants, 155 complexes, 154–156 stability constants of complexes, 155 structure, 151 Uridine 5⬘-triphosphate, see 5⬘-UTP

SUBJECT INDEX

701

Urinary tract infections, 242 5⬘-UTP, 146, 512 complexes, 141, 143 UV-Vis spectrophotometry (of) acetyl-coenzyme A synthase model, 209, 214 acireductone dioxygenase, 486, 489, 491 F430, 329–332, 338 methyl-coenzyme M reductase, 338–340, 342 [NiFe] hydrogenases, 192, 532 nickel superoxide dismutase, 429, 432 urease models, 222

V Vaccination against Helicobacter pylori infection, 568, 569 Vaccinium myrtillus, 18 vitis-idaea, 18 Vanadate and urease, 267 Vasopressin (complex with) arginine8-, 85 Cu(II), 85 Ni(II), 78 Vasotocin arginine8-, 85 Cu(II) complex, 85 Vertebrate (see also individual names) sperm, 84 Virus (see also individual names) Coxsackie, 630 hepatitis C, 481, 483 tobacco mosaic, 448 turnip mosaic, 49 Vitamin(s) (see also individual names) B2, 148 C, see Ascorbate Voltammetry adsorptive cathodic stripping, 5 cyclic, see Cyclic voltammetry nickel samples, 5

[Voltammetry] [NiFe] hydrogenases, 306, 307 protein film, 306, 307

W Waste municipal, 41, 42 nickel in, 2, 7, 41–43 Water lake, see Lake nickel concentrations, 10–12, 41, 42 nickel contaminated, 622, 623 sea-, 110 Wheat nickel toxicity, 18 Wolinella succinogenes, 559 Wood-Ljungdahl pathway, 390, 391

X XAFS nickel samples, 5 XANES acireductone dioxygenase, 474, 486, 487 glyoxalase I, 455 methyl-coenzyme M reductase, 339 nickel K-edge, 427, 487 nickel samples, 5 nickel superoxide dismutase, 426–428 [NiFe] hydrogenases, 299 sulfur K-edge, 428 Xanthosine 5⬘-monophosphate, see 5⬘-XMP XAS, see X-ray absorption spectroscopy Xenobiotics, 598 Xenon probe, 293, 294 Xenopus laevis, 632 5⬘-XMP acidity constants, 146 complexes, 146–148 structure, 147, 148

702

X-ray absorption fine-structure spectroscopy, see XAFS X-ray absorption near-edge structure spectroscopy, see XANES X-ray absorption spectroscopy (studies of), 47, 50 acetyl-coenzyme A synthase, 204 acireductone dioxygenase, 474, 480, 483, 490 iron K-edge, 378 methyl-coenzyme M reductase, 339, 341, 342 near-edge, 187 Ni(I) complexes, 187 nickel L-edge, 299, 378 nickel K-edge, 299, 365, 378, 426 nickel superoxide dismutases, 422, 426–428 [NiFe] hydrogenase (models), 192, 199, 282, 299, 300, 306, 310 sulfur K-edge, 422, 426–428 urease, 267 X-ray crystal structure studies of (see also Crystal structures) acetyl-coenzyme A synthase models, 206, 209 acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 362, 525 carbon monoxide dehydrogenase, 200 Cu(Cyd)2⫹ complex, 123 methyl-coenzyme M reductase, 525 [NiFe] hydrogenases, 291–294 Ni(II)-peptide complexes, 89 Ni(II)-purine complexes, 118, 119, 145 Ni(II)-pyridine complexes, 126 Ni(II)-sugar complexes, 113–115 urease, 256, 259, 521 X-ray diffraction spectroscopy studies of nickel samples, 5 [NiFe] hydrogenases, 192 X-ray fluorescence spectrometry (studies of) nickel samples, 5 synchrotron-based, 5

SUBJECT INDEX

Y Yeast (see also Saccharomyces cerevisiae), 529, 587, 645 acireductone dioxygenase, 475 copper metallochaperone, 531 nickel uptake, 38 Yersinia sp., 560 pestis, 451, 453, 460–462

Z Zinc (different oxidation states) accumulating plants, see Hyperaccumulating plants interdependency with other metal ions, see Interdependencies transporter, 38, 46 Zinc(II) (in/complexes with) acetyl-coenzyme A synthases/carbon monoxide dehydrogenases, 374, 376 amino acids, 95 carbonyl group, 117 glyoxalase I, see Glyoxalase I histidine, 95 hormones, see Hormones hydroxyl group, 117 nucleic acid binding, 165, 167 nucleotides, 137, 142, 144, 148, 150, 157 peptide, see Peptides phosphate, 129, 130 purine-nucleotide (N1)-oxide, 153 SlyD, 512 ternary ATP-buffer complexes, 163 ternary nucleotide complexes, 164 thiophosphate, 155, 156 urease, 266, 267 water exchange rate, 111 Zinc finger proteins, 81, 83 transcription factor, 597

Nickel and Its Surprising Impact in Nature: Metal Ions in Life Sciences (Vol. 2)

Neurodegenerative Diseases and Metal Ions: Metal Ions in Life Sciences Vol 1

Metal Ions in Fungi

Organometallics in Environment and Toxicology (Metal Ions in Life Sciences, Volume 7)

Biosorbents for Metal Ions

Metallothioneins and Related Chelators (Volume 5) (Metal Ions in Life Sciences)

Metal Ions and Neurodegenerative Disorders

Structural Proteomics And Its Impact On The Life Sciences

The Ubiquitous Roles of Cytochrome P450 Proteins (Metal Ions in Life Sciences, Volume 3)

Energy And Life (Modules in Life Sciences)

Interplay between Metal Ions and Nucleic Acids

Life Itself: Its Origin and Nature

Life Itself: Its Origin and Nature

Knowledge and Its Place in Nature

Information and its role in nature

Improvisation: Its Nature and Practice In Music

Ions in Polymers

Philosophy of Nature, Vol. 2

Forest Life vol 2

Encyclopaedia of Biological Disaster Management: vol. 2. Biological Disaster: Its Impact on Health and its Management

Vol 13 Metal Carbenes in Organic Synthesis

Brilliant Light in Life and Material Sciences

Bioinformatics: applications in life and environmental sciences

Visualization in medicine and life sciences II

Meditation and Its Utility in Daily Life

Colloids and Interfaces in Life Sciences

Nickel

Ionization in Gases by Ions and Atoms

Ultratrace Metal Analysis in Biological Sciences and Environment

Electrons and Ions in Liquid Helium

Timber: Its Nature and Behaviour

Nickel and Its Surprising Impact in Nature: Metal Ions in Life Sciences (Vol. 2)

Neurodegenerative Diseases and Metal Ions: Metal Ions in Life Sciences Vol 1

Metal Ions in Fungi

Organometallics in Environment and Toxicology (Metal Ions in Life Sciences, Volume 7)

Biosorbents for Metal Ions

Metallothioneins and Related Chelators (Volume 5) (Metal Ions in Life Sciences)

Metal Ions and Neurodegenerative Disorders

Structural Proteomics And Its Impact On The Life Sciences

The Ubiquitous Roles of Cytochrome P450 Proteins (Metal Ions in Life Sciences, Volume 3)

Energy And Life (Modules in Life Sciences)

Interplay between Metal Ions and Nucleic Acids

Life Itself: Its Origin and Nature

Life Itself: Its Origin and Nature

Knowledge and Its Place in Nature

Information and its role in nature

Improvisation: Its Nature and Practice In Music

Ions in Polymers

Philosophy of Nature, Vol. 2

Forest Life vol 2

Encyclopaedia of Biological Disaster Management: vol. 2. Biological Disaster: Its Impact on Health and its Management

Vol 13 Metal Carbenes in Organic Synthesis

Brilliant Light in Life and Material Sciences

Bioinformatics: applications in life and environmental sciences

Visualization in medicine and life sciences II

Meditation and Its Utility in Daily Life

Colloids and Interfaces in Life Sciences

Nickel

Ionization in Gases by Ions and Atoms

Ultratrace Metal Analysis in Biological Sciences and Environment

Electrons and Ions in Liquid Helium

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