Modern Biooxidation: Enzymes, Reactions and Applications

Modern Biooxidation Edited by Rolf D. Schmid and Vlada B. Urlacher 1807–2007 Knowledge for Generations Each generatio...

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Modern Biooxidation Edited by Rolf D. Schmid and Vlada B. Urlacher

1807–2007 Knowledge for Generations Each generation has its unique needs and aspirations. When Charles Wiley first opened his small printing shop in lower Manhattan in 1807, it was a generation of boundless potential searching for an identity. And we were there, helping to define a new American literary tradition. Over half a century later, in the midst of the Second Industrial Revolution, it was a generation focused on building the future. Once again, we were there, supplying the critical scientific, technical, and engineering knowledge that helped frame the world. Throughout the 20th Century, and into the new millennium, nations began to reach out beyond their own borders and a new international community was born. Wiley was there, expanding its operations around the world to enable a global exchange of ideas, opinions, and know-how. For 200 years, Wiley has been an integral part of each generation’s journey, enabling the flow of information and understanding necessary to meet their needs and fulfill their aspiration. Today, bold new technologies are changing the way we live and learn. Wiley will be there, providing you the must-have knowledge you need to imagine new worlds, new possibilities, and new opportunities. Generations come and go, but you can always count on Wiley to provide you the knowledge you need, when and where you need it!

William J. Pesce President and Chief Executive Officer

Peter Booth Wiley Chairman of the Board

Modern Biooxidation Enzymes, Reactions and Applications Edited by Rolf D. Schmid and Vlada B. Urlacher

The Editors Prof. Dr. Rolf D. Schmid Universität Stuttgart Institut für Technische Biochemie Allmandring 31 70569 Stuttgart Germany

All books published by Wiley-VCH are carefully produced. Nevertheless, authors, editors, and publisher do not warrant the information contained in these books, including this book, to be free of errors. Readers are advised to keep in mind that statements, data, illustrations, procedural details or other items may inadvertently be inaccurate.

Dr. Vlada B. Urlacher-Kursif Universität Stuttgart Institut für Technische Biochemie Allmandring 31 70569 Stuttgart Germany

Library of Congress Card No.: applied for

Cover Crystal structure of human cytochrome p450 2C9 (PDB entry 1OG5A) complexed with substrate warfarin (green) [1]. The mesh represents the substrate binding cavity inside the enzyme. The approach of the substrate (blue) to the heme (red) was observed during molecular dynamics simulations [2].

Bibliographic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliografie; detailed bibliographic data are available in the Internet at 〈http://dnb.d-nb.de〉.

1 Williams, P.A., Cosme, J., Ward, A., Angove, H.C., Matak Vinkovic, D., Jhoti, H. Crystal structure of human cytochrome p450 2C9 with bound warfarin. Nature 2003, 424, 464–468. 2 Seifert, A., Tatzel, S., Schmid, R.D., Pleiss, J. Multiple molecular dynamics simulations of human p450 monooxygenase CYP2C9: the molecular basis of substrate binding and regioselectivity toward warfarin. Proteins 2006, 64(1), 147–155.

British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library.

© 2007 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim All rights reserved (including those of translation into other languages). No part of this book may be reproduced in any form – by photoprinting, microfi lm, or any other means – nor transmitted or translated into a machine language without written permission from the publishers. Registered names, trademarks, etc. used in this book, even when not specifically marked as such, are not to be considered unprotected by law. Composition SNP Best-set Typesetter Ltd., Hong Kong Printing Betz-Druck GmbH, Darmstadt Bookbinding Litges & Dopf GmbH, Heppenheim Cover Design Schulz Grafi k-Design, Fußgönheim Wiley Bicentennial Logo Richard J. Pacifico Printed in the Federal Republic of Germany Printed on acid-free paper ISBN 978-3-527-31507-9

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Contents

1

1.1 1.2 1.2.1 1.2.2 1.2.3 1.3 1.3.1 1.3.1.1 1.3.1.2 1.3.1.3 1.3.2 1.3.2.1 1.3.2.2 1.3.2.3 1.3.3 1.3.3.1 1.3.3.2 1.3.3.3 1.3.3.4 1.3.3.5 1.3.3.6 1.3.3.7 1.3.3.8 1.3.4 1.4 1.4.1

Biooxidation with PQQ- and FAD-Dependent Dehydrogenases 1 Osao Adachi, Yoshitaka Ano, Hirohide Toyama, and Kazunobu Matsushita Introduction 1 Basic Technical Information Regarding Membrane-bound Enzymes 4 Preparation of Cytosolic Fractions and Membrane Fractions 4 EDTA Treatment of the Membrane Fraction Carrying PQQ as Coenzyme 5 Assays of Enzyme Activity 5 PQQ-Dependent Dehydrogenases 6 Alcohol Oxidation 6 Membrane-Bound Alcohol Dehydrogenase (ADH III) 6 Soluble Alcohol Dehydrogenases 9 Cyclic Alcohol Dehydrogenase (Secondary Alcohol Dehydrogenase), Membrane-Bound 9 Glucose Oxidation 11 Membrane-Bound D -Glucose Dehydrogenase (m-GDH) 11 Soluble D -Glucose Dehydrogenase (s-GDH) 12 Applications of Quinoprotein GDHs as D -Glucose Sensors 13 Polyol Oxidation 14 D -Arabitol Dehydrogenase, Membrane-Bound 14 meso-Erythritol Oxidation Dehydrogenase, Membrane-Bound 16 D -Gluconate Oxidizing Polyol Dehydrogenase, Membrane-Bound 17 Glycerol Dehydrogenase, Membrane-Bound 19 D -Mannitol Dehydrogenase, Membrane-Bound 20 Ribitol Dehydrogenase, Membrane-Bound 21 D -Sorbitol Dehydrogenase, Membrane-Bound 22 L-Sorbosone Dehydrogenase, Membrane-Bound 23 Quinate Oxidation. Membrane-Bound Quinate Dehydrogenase (QDH) 24 FAD-Dependent Dehydrogenase 27 D -Fructose Dehydrogenase, Membrane-Bound 27

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Contents

1.4.2 1.4.3 1.4.4 1.4.5 1.5 1.5.1

Dehydrogenase, Membrane-Bound 28 Dehydrogenase, Membrane-Bound 29 2-Keto- D -gluconate Dehydrogenase, Membrane-Bound 31 Sorbitol Dehydrogenase, Membrane-Bound 32 Miscellaneous 33 Aldehyde Dehydrogenase, Membrane-Bound 33 References 35

2

Catalytic Applications of Laccase 43 Feng Xu, Ture Damhus, Steffen Danielsen, and Lars Henrik Østergaard Properties of Classical Laccase 43 Structure 43 Enzymology 44 4As Industrial Catalysts 46 Advantages 46 Shortcomings 48 Applications of Laccase for Industrial Oxidation Processes 48 Laboratory-Level Trials 49 Delignification 49 Dye and Colorant Bleaching 50 Bioremediation 50 Other Degradation Applications 51 Functional Biotransformation 51 Biosensing 53 Desirable Application Modes 53 Commercialized Applications 55 Preventing Taint in Cork Stoppers 56 Denim Bleaching 56 Paper Mill Effluent Treatment and Cardboard Strengthening 56 Major Hurdles to Further Development from Laboratory Trials 57 More Recent Developments 57 Novel Laccase Catalytic Systems 57 New Laccases 57 New Mediators 60 Cooperation with Other Enzymes 62 New Leads for Laccase Application 62 Laccase-Based Defense Against Biological and Chemical Warfare Agents 62 Degradation of PAH, Plastics, or Lipids 63 Enzymatic Fuel Cells/Batteries 64 Novel Synthetic Applications 65 Biorefinery 66 Further Developing Laccase Catalysis 66 Laccase Engineering 66

2.1 2.1.1 2.1.2 2.1.3 2.1.3.1 2.1.3.2 2.2 2.2.1 2.2.1.1 2.2.1.2 2.2.1.3 2.2.1.4 2.2.1.5 2.2.1.6 2.2.1.7 2.2.2 2.2.2.1 2.2.2.2 2.2.2.3 2.2.2.4 2.3 2.3.1 2.3.1.1 2.3.1.2 2.3.1.3 2.3.2 2.3.2.1 2.3.2.2 2.3.2.3 2.3.2.4 2.3.2.5 2.4 2.4.1

D -Gluconate

D -Hexosamine

Contents

2.4.2

Laccase Production References 68

67

3

Biocatalytic Scope of Baeyer–Villiger Monooxygenases 77 Marco W. Fraaije and Dick B. Janssen 3.1 Introduction 77 3.1.1 The Baeyer–Villiger Reaction 77 3.1.2 Baeyer–Villiger Biocatalysts: Classification and Occurrence 78 3.1.2.1 Type I Baeyer–Villiger Monooxygenases 78 3.1.2.2 Type II Baeyer–Villiger Monooxygenases 78 3.1.2.3 Alternative Baeyer–Villiger Biocatalysts 79 3.2 Type I Baeyer–Villiger Monooxygenases: Versatile Oxidative Biocatalysts 80 3.2.1 Mechanistic and Structural Properties of Type I BVMOs 80 3.2.2 Diversity 84 3.2.3 Molecular Features 86 3.2.4 Kinetic Characteristics 86 3.2.5 Coenzyme Dependency 87 3.2.6 Uncoupling and Overoxidation 88 3.2.7 Biocatalyst Stability 88 3.2.8 Substrate Specificity 89 3.2.9 Unexplored Type I BVMOs 90 3.2.10 Mining Genomes for Novel BVMOs 92 3.3 Concluding Remarks 93 References 94 4

4.1 4.2 4.3 4.3.1 4.3.2 4.3.3 4.3.4 4.4

5

5.1 5.2

The Bacterial Cytochrome P450 Monooxygenases: P450cam and P450BM-3 99 Vlada B. Urlacher, Stephen G. Bell, and Luet-Lok Wong Introduction 99 Biotransformation by Bacterial P450 Enzymes 99 General Features of P450cam and P450BM-3 102 Aromatic Compounds 105 Alkanes and Alicyclics 109 Terpenoid Compounds 111 Human Metabolites 114 The Scope of P450 Engineering 116 References 117 Cytochrome P450 Redox Partner Systems: Biodiversity and Biotechnological Implications 123 Andrew W. Munro, Hazel M. Girvan, Joseph P. McVey, and Kirsty J. McLean Introduction 123 P450 Redox Partners 124

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5.2.1 5.2.2 5.2.3 5.2.4 5.2.5 5.2.6 5.2.7 5.3 5.4 5.5 5.6 5.7

6

6.1 6.2 6.3 6.3.1 6.3.2 6.3.3 6.3.4 6.4 6.4.1 6.4.2 6.5

7

7.1 7.2 7.2.1 7.2.1.1 7.2.1.2 7.2.2 7.2.3 7.2.3.1

A “Historical” Perspective 124 The P450 Catalytic Cycle and Electron Transfer Events 125 P450cam and its Reductase System 127 Adrenodoxin and Adrenodoxin Reductase 128 Cytochrome P450 Reductase 129 P450BM-3 and Related CPR Fusion Enzymes 131 A Novel Class of P450–Redox Partner Fusion Enzymes 136 Increasing P450–Redox Partner Complexity: Flavodoxins and Diverse Ferredoxins 137 Natural and Artificial P450–Redox Partner Fusion Enzymes and their Biocatalytic Potential 138 Other Routes to Driving P450 Catalytic Function 140 Uncoupling, Enzyme Stability and Coenzyme Issues 142 Future Prospects 143 References 145 Steroid Hydroxylation: Microbial Steroid Biotransformations Using Cytochrome P450 Enzymes 155 Matthias Bureik and Rita Bernhardt Introduction 155 Cytochrome P450-Dependent Steroid Hydroxylase Systems 156 Native Microorganisms in Steroid Biotransformation 159 11α-Hydroxylation 160 11β-Hydroxylation 161 16α-Hydroxylation 162 Conclusions 163 Genetically Modified Microorganisms in Steroid Biotransformation 163 Soluble Cytochromes P450 164 Membrane-Bound Cytochromes P450 166 Synopsis and Concluding Remarks 170 References 171 A Modular Approach to Biotransformation Using Microbial Cytochrome P450 Monooxygenases 177 Akira Arisawa and Hitosi Agematu Introduction 177 Experimental Outline 180 Gene Sequences 180 pT7NS-camAB 180 Plasmids to Express Bacterial CYPs 180 Preparation of Whole Cell Catalysts 181 Biotransformation of the CYP Substrates 181 Carbomycin A 181

Contents

7.2.3.2 7.2.3.3 7.2.4 7.3 7.4 7.5 7.6

Pravastatin 182 7-Hydroxycoumarin 182 Biotransformation by CYP Reaction Array 182 Bacterial CYP Expression System in E. coli 183 Construction of a Bacterial CYP Library 185 Construction of a Bacterial CYP Reaction Array 186 Application of the CYP Reaction Array to Biotransformation Screening 187 References 190

8

Selective Microbial Oxidations in Industry: Oxidations of Alkanes, Fatty Acids, Heterocyclic Compounds, Aromatic Compounds and Glycerol Using Native or Recombinant Microorganisms 193 Albrecht Weiss 8.1 Introduction 193 8.2 Selective Oxidation of Hydrocarbons and Fatty Acids 194 8.2.1 Alkane Oxidation to Medium-Chain Alcohols [11] 194 8.2.2 Alkane and Fatty Acid Oxidation to Dicarboxylic Acids 196 8.2.2.1 Alkanes 197 8.2.2.2 Dicarboxylic Acids 197 8.3 Aromatic Compounds/Fine Chemicals 198 8.3.1 Conversion of Toxic Compounds: Catechols 198 8.3.2 Production of (R)-2-(4-Hydroxyphenoxy)propionic Acid 199 8.3.3 Selective Oxidation to Aromatic Aldehydes with Recombinant Cells 200 8.3.4 Styrene Oxide Production in a Two-Liquid Phase System 200 8.4 Heterocyclic Compounds 200 8.4.1 Enzymatic Oxidation of Methyl Groups in Aromatic Heterocycles 201 8.4.2 Preparation of 6-Hydroxynicotinic Acid 202 8.4.3 Preparation of 5-Hydroxypyrazinecarboxylic Acid 202 8.4.4 Preparation of 6-Hydroxy-(S)-nicotine and 4-[6-Hydroxypyridin-3-yl]4-oxobutyrate 202 8.4.5 Bulk Chemicals/Indigo 203 8.5 Glycerol Conversion to Dihydroxyacetone 206 8.6 Perspectives 207 References 207 9

9.1 9.2 9.3

Preparation of Drug Metabolites using Fungal and Bacterial Strains 211 Oreste Ghisalba and Matthias Kittelmann Introduction 211 Phase I Drug-Metabolizing Enzymes 212 Needs and “Platforms” for the Generation of Drug Metabolites 214

IX

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Contents

9.3.1 9.3.2 9.4 9.4.1 9.4.2 9.5 9.6 9.7 9.7.1 9.7.2

9.8 9.8.1 9.9

10

10.1 10.1.1 10.1.2 10.1.3 10.2 10.3 10.3.1 10.3.2 10.3.2.1 10.3.2.2 10.3.2.3 10.3.2.4 10.3.3 10.3.3.1 10.3.3.2 10.3.3.3 10.3.3.4

Recombinant Human Cytochrome P450 (rhCYP) Systems (acquired from British Technology Group/University of Dundee) 215 Microbial Strains Performing Oxidative Reactions (in-house technology) 215 Microbial Models for Oxidative Drug Metabolism 215 2Prokaryotic P450s 218 Microbial Eukaryotic P450s 218 Correlation of Microbial and Mammalian Oxidative Drug Metabolism 221 Correlation of Microbial Reactions with Human CYP Isozyme-Specific Reactions 221 Novartis Research Examples of Microbial Hydroxylations 225 Preparation of 10,11-Epoxy-carbamazepine and 10,11-Dihydro-10-hydroxy-carbamazepine 225 Preparation of 4-(4′-Hydroxyanilino)-5-anilinophthalimide and 4,5-Bis-(4′-hydroxyanilino)-phthalimide by Microbial Hydroxylation 227 Microbial Oxidation of Natural Products 228 Microbial Hydroxylation and Epoxidation of Milbemycins 229 Conclusions 229 References 231 Recombinant Yeast and Bacteria that Express Human P450s: Bioreactors for Drug Discovery, Development, and Biotechnology 233 Steven P. Hanlon, Thomas Friedberg, C. Roland Wolf, Oreste Ghisalba, and Matthias Kittelmann Background 234 Importance of Recombinant P450s for Drug Development 234 Fundamentals of Heterologous Expression in Bacteria 235 Fundamentals of Heterologous Expression in Yeast 236 Comparison of P450 Levels and Enzymic Activities in Various Models 237 Use of E. coli P450 Expression Systems in Bioreactors 240 General Considerations 240 The Roche Experience 240 Background and Utility of P450 Systems in Pharma Research 240 Fermentation of Recombinant E. coli 241 Biotransformations Catalyzed by Recombinant CYP450 241 Preparation of N-Desethyl Amodiaquine 242 The Novartis Experience 244 Introduction 244 Production of E. coli Cells with CYP Activity 244 Whole Cell Biotransformation 246 Recent Developments 246

Contents

10.4

Conclusion 246 References 247

11

Human Cytochrome P450 Monooxygenases – a General Model of Substrate Specificity and Regioselectivity 253 Jürgen Pleiss Introduction 253 What Can We Learn From Sequence? 254 The Cytochrome P450 Engineering Database (CYPED) 254 The Effect of Mutations on Activity 255 What Can We Learn from Structure? 258 2The Role of Flexibility 258 The Role of Binding Site Shape 259 Conclusion 261 References 262

11.1 11.2 11.2.1 11.2.2 11.3 11.3.1 11.3.2 11.4

12

12.1 12.2 12.3 12.4 12.5 12.5.1 12.5.2 12.5.3 12.5.4 12.5.5 12.5.6 12.6 12.7 12.7.1 12.7.2 12.7.3 12.7.4 12.8

Approaches to Recycling and Substituting NAD(P)H as a CYP Cofactor 265 Dirk Holtmann and Jens Schrader Introduction 265 Chemical Substitution of Cofactors 266 Enzymatic Regeneration of Cofactors 267 Photochemical Approaches to Substituting or Regenerating Cofactors for P450 Systems 271 Electrochemical Systems for Substitution or Regeneration of Cofactors 272 Electrochemical Regeneration of Natural Cofactors 273 Electrochemical Regeneration of Artificial Cofactors 274 Electrochemical Generation of Hydrogen Peroxide 275 Electrochemistry of P450 at Modified Electrodes 275 Electrochemistry of P450 in Surfactant Films 276 Incorporation of Cytochrome P450 in Conducting Polymers 278 Redox Mediators 278 Molecular Biological Approaches 280 Peroxide Shunt 280 Artificial Electron Transfer Systems 281 Changing the Cofactor Specificity of P450 Systems 281 Intracellular Cofactor Regeneration 282 Conclusion and Outlook 282 References 284 Index

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XIII

Preface

In recent decades, biochemical science and technology has made tremendous progress. A recent survey among US universities has revealed that nearly half of academic funding is devoted to the life sciences [1], and a similar situation prevails in most other industrialized countries. In spite of this massive investment, the share of biotechnology in the production of energy and chemicals is small – educated estimates are in the range of 3% (R Diercks, BASF AG, personal communication). Thus, much remains to be done to develop the “biorefinery” concept (see, e.g. ref. [2]) from vision into reality. To date, enzyme technology in industry has been mostly restricted to selective hydrolyses or ester/amide bond formations. Important unit operations in chemical synthesis such as C-C bond formation or selective oxidations have remained the domain of the synthetic organic chemist, who can find in his or her textbooks a vast number of protocols for the transformation of many, if not most types of possible structures. Enzymatic methods have been less successful in these domains, major stumbling blocks being the lack of enzymes with the required selectivity and the need for expensive cofactors such as NADH or ATP. More recently, techniques such as protein engineering, directed evolution, cofactor regeneration, and metabolic engineering have opened up new avenues to remove these bottlenecks. In addition, new types of enzymes have appeared on the stage of enzyme technology. Thus, among the oxidizing enzymes cytochrome and flavine monooxygenases, a class of enzymes hitherto unknown in the context of enzyme technology, are now being hotly investigated for use in fine chemical selective oxidation. Also laccases, mostly known for their potential as bleaching agents for paper pulp, are just being rediscovered as candidates for selective biooxidation. Research in advanced enzyme technology is global. In fact, a Japanese program on “Green Biotechnology”, inaugurated as early as 2000 and now in its second 5year cycle (S Shimizu, Coordinator of METI project “Green Biotechnology”, personal communication), has made significant advances towards the design of selective oxidative enzymes. It is thus our great privilege to have collected 12 contributions from leaders in this area of research with pharmaceutical or chemical backgrounds, from

XIV

Preface

industry and from academia, working in Europe, the USA and Japan, to provide insight into the state of the art in work to turn oxidizing enzymes into a tool for the pharmaceutical and chemical industries. At this point we wish to acknowledge the excellent cooperation of the authors of this book, who all submitted manuscripts and corrections on time and without any delay. Thanks are also due to Dr Waltraud Wuest and colleagues at Wiley-VCH Publishers in Weinheim, Germany, who did a proficient job in editing the manuscript, and to Mr Alexander Seifert of our institute who provided us with a fi ne graph for the cover. Finally, we are grateful to the German Research Foundation (DFG) for financial support of the Cooperative Research Center “Selective oxidation using molecular oxygen” (SFB706) and the Ministry of Science, Research and the Arts BadenWuerttemberg for institutional funding. Stuttgart, Summer 2007

Professor Dr. Rolf D. Schmid Dr. Vlada B. Urlacher

References 1 Rovner, S.L. Academic R&D spending trends. Chem Eng News 2007, 84:51, 47–51. 2 Ritter, S. Biorefi nery gels ready to deliver the goods. Chem Eng News 2006, 84:34, 47.

XV

List of Contributors Osao Adachi Yamaguchi University Department of Biological Chemistry Yamaguchi 753-8515 Japan Hitosi Agematu Akita National College of Technology Department of Applied Chemistry 1-1 Iijima-Bunkyo-cho, Akita 110-8511 Japan Yoshitaka Ano Yamaguchi University Department of Biological Chemistry Yamaguchi 753-8515 Japan Akira Arisawa Mercian Corporation Bioresource Laboratories 1808 Nkaizumi, Iwata, Shizuoka 438-0078 Japan

Stephen G. Bell University of Oxford Department of Chemistry Inorganic Chemistry Laboratory South Parks Road Oxford OX1 3QR UK Rita Bernhardt Saarland University Institute of Biochemistry P.O. Box 151150 66123 Saarbrücken Germany Matthias Bureik Saarland University Institute of Biochemistry P.O. Box 151150 66123 Saarbrücken Germany Ture Damhus Novozymes A/S Krogshoejvej 36 2880 Bagsvaerd Denmark Steffen Danielsen Novozymes A/S Krogshoejvej 36 2880 Bagsvaerd Denmark

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Contributors

Thomas Friedberg University of Dundee Ninewells Hospital and Medical School Biomedical Research Centre Dundee DD1 9SY UK

Dirk Holtmann DECHEMA e.V. Karl-Winnacker-Institute Biochemical Engineering Group Theodor-Heuss-Allee 25 60486 Frankfurt am Main Germany

Marco W. Fraaije University of Groningen Groningen Biomolecular Sciences and Biotechnology Institute Biochemical Laboratory Nijenborgh 4 9747 AG Groningen The Netherlands

Dick B. Janssen University of Groningen Groningen Biomolecular Sciences and Biotechnology Institute Biochemical Laboratory Nijenborgh 4 9747 AG Groningen The Netherlands

Oreste Ghisalba Novartis Pharma AG Novartis Institutes for BioMedical Research Discovery Technologies, WSJ-508.202 A 4002 Basel Switzerland

Matthias Kittelmann Novartis Pharma AG Novartis Institutes for Biomedical Research Discovery Technologies, WSJ-508.202 A 4002 Basel Switzerland

Hazel M. Girvan University of Manchester Manchester Interdisciplinary Biocentre, SCEAS 131 Princess Street Manchester M1 7DN UK

Kazunobu Matsushita Yamaguchi University Department of Biological Chemistry Yamaguchi 753-8515 Japan

Steven P. Hanlon F. Hoffmann-La Roche Ltd Safety & Technical Sciences Basel Department of Chemical Synthesis, Biocatalysis Bldg 62/136 4070 Basel Switzerland

Kirsty J. McLean University of Manchester Manchester Interdisciplinary Biocentre, SCEAS 131 Princess Street Manchester M1 7DN UK

Contributors

Joseph P. McVey University of Manchester Manchester Interdisciplinary Biocentre, SCEAS 131 Princess Street Manchester M1 7DN UK

Vlada B. Urlacher University of Stuttgart Institute of Technical Biochemistry Allmandring 31 70569 Stuttgart Germany

Andrew W. Munro University of Manchester Manchester Interdisciplinary Biocentre, SCEAS 131 Princess Street Manchester M1 7DN UK

Albrecht Weiss Cognis Deutschland GmbH CRT-Z 43 Postfach 130164 40551 Düsseldorf Germany

Lars Henrik Østergaard Novozymes A/S Krogshoejvej 36 2880 Bagsvaerd Denmark Jürgen Pleiss University of Stuttgart Institute of Technical Biochemistry Allmandring 31 70569 Stuttgart Germany Jens Schrader DECHEMA e.V. Karl-Winnacker-Institute Biochemical Engineering Group Theodor-Heuss-Allee 25 60486 Frankfurt am Main Germany Hirohide Toyama Yamaguchi University Department of Biological Chemistry Yamaguchi 753-8515 Japan

C. Roland Wolf University of Dundee Ninewells Hospital and Medical School Biomedical Research Centre Dundee DD1 9SY UK Luet-Lok Wong University of Oxford Department of Chemistry Inorganic Chemistry Laboratory South Parks Road Oxford OX1 3QR UK Feng Xu Novozymes, Inc. 1445 Drew Avenue Davis, CA 95616 USA

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1 Biooxidation with PQQ- and FAD-Dependent Dehydrogenases Osao Adachi, Yoshitaka Ano, Hirohide Toyama, and Kazunobu Matsushita

Summary

Among the obligate aerobic bacteria, acetic acid bacteria are well known for their powerful ability to oxidize alcohols, sugars, or sugar alcohols and to accumulate the corresponding oxidation products in the culture medium. These reactions are restricted to one-step incomplete oxidation (so-called oxidative fermentation) and are catalyzed by primary dehydrogenases located on the outer surface of the cytoplasmic membrane, the active sites of which face the periplasmic space. All enzyme activities are linked, without exception, to the terminal ubiquinol oxidase via ubiquinone in the respiratory chain of the organisms. The respective primary dehydrogenases working in the periplasmic sugar and alcohol respirations include many unique pyrroloquinoline quinone (PQQ)-dependent dehydrogenases (quinoproteins and quinoprotein–cytochrome c complexes) and flavin adenine dinucleotide (FAD)-dependent dehydrogenases (flavoprotein–cytochrome c complexes). Since this sugar and alcohol respiration does not seem to generate much energy, acetic acid bacteria use rapid oxidation to produce a large number of oxidation products, compensating for the necessary bioenergy required. We have learnt a lot about the biological activities of acetic acid bacteria in the past hundred years. Among them are classic but typically important microbial bioconversions for practical use, such as the production of vinegar, D -gluconate, and L-sorbose. However, our understanding of the molecular mechanisms remains to be clarified. We have been trying to uncover the enzymatic and biochemical mechanisms of the respective enzymes in acetic acid bacteria since the 1970s. In this chapter, the properties and characteristics of the individual enzymes involved in oxidative fermentation are exemplified. 1.1 Introduction

Many membrane-bound dehydrogenases in the periplasmic space or on the outer surface of the cytoplasmic membrane of acetic acid bacteria and other aerobic

2

1 Biooxidation with PQQ- and FAD-Dependent Dehydrogenases

Gram-negative bacteria have been classified as PQQ- or FAD-dependent dehydrogenases [1]. Most of the enzymes are closely associated with oxidative fermentation in industry, catalyzing an incomplete one-step oxidation, allowing accumulation of an equivalent amount of corresponding oxidation products outside the cells. The active sites of individual enzymes face the periplasmic space as illustrated in Fig. 1.1. All enzyme reactions are carried out by periplasmic oxidase systems, including alcohol- and sugar-oxidizing enzymes of the organisms. D -Glucose, ethanol, and many other substrates are oxidized by the dehydrogenases (shown as PQQ or FAD, except for aldehyde dehydrogenase) that are tightly bound to the outer surface of the cytoplasmic membranes of the organism. These membrane-bound enzymes irreversibly catalyze incomplete one-step oxidation and the corresponding oxidation products accumulate rapidly in the culture medium or reaction mixture. The electrons (e− ) generated by the action of these dehydrogenases are transferred to ubiquinone in the membrane. The reducing equivalents are further transferred to the terminal ubiquinol oxidase in the cytoplasmic membranes. Thus, the organisms generate bioenergy through the enzyme activities of PQQand FAD-dependent dehydrogenases. The outer membrane of the organism forming the periplasmic space is omitted from Fig. 1.1. Many different NAD- or NADP-dependent dehydrogenases in the cytoplasm have no function in oxidative fermentation, and thus are not shown in Fig. 1.1.

Fig. 1.1 Membrane-bound PQQ- and FAD-dependent primary dehydrogenases on the outer surface of acetic acid bacteria.

1.1 Introduction

At present, of the enzymes exploited as either PQQ-dependent or FADdependent dehydrogenases, aldehyde dehydrogenase is the only one that is known to use a molybdopterin coenzyme. Unlike the cytoplasmic oxidoreductases, no energy is required for substrate intake into the periplasm and pumping out the oxidation products across the outer membrane as shown in Fig. 1.2. Microbial production of L-sorbose, aldehyde, and 2-keto- D -gluconate are the examples shown in Fig. 1.2. All substrates are oxidized by the respective membrane-bound dehydrogenase, of which active site faces the periplasmic space formed between the outer membrane and the cytoplasmic membrane. The dehydrogenase then donates electrons to ubiquinone (UQ) that in turn transfers them to the terminal ubiquinol oxidase. The terminal oxidase generates an electrochemical proton gradient either by charge separation or by a proton pump or by both during substrate oxidation by the membrane-bound enzymes, allowing the organism to acquire bioenergy through substrate oxidation. Traditionally, acetate fermentation (vinegar production) and L-sorbose fermentation are typical examples of oxidative fermentation and the classic case of microbial bioconversion. Our understanding of the mechanism of oxidative fermentation, however, was not elucidated until relatively recently. Most enzymes involved in oxidative fermentation are associated closely with industrial applications for useful biomaterial production. The production of acetate, L-sorbose, D gluconate, dihydroxyacetone, and others developed as a practical industry before the clarification of the molecular mechanisms of the responsible enzymes. It was in 1970s that we started to clarify the molecular mechanisms of the individual enzymes involved in oxidative fermentation. Before describing the actions of the individual PQQ- and FAD-dependent dehydrogenases, it is worth clarifying the common physiological roles and localizations of PQQ- and FAD-dependent dehydrogenases in acetic acid bacteria and

Fig. 1.2 Membrane-bound dehydrogenase-dependent periplasmic oxidase systems.

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other microorganisms. Many people still believe that acetate is produced by the cytosolic NAD(P)-dependent alcohol dehydrogenase and keto- D -gluconate by the cytosolic NADP-dependent D -gluconate dehydrogenase located in the cytoplasm. Such a serious confusion is probably caused by the confused of localization of the enzymes concerned. Both membrane-bound enzymes and NAD(P)-dependent enzymes sometimes occur in the same cell-free extract when bacterial cells are broken down and the cell-free extract is prepared. Some periplasmic enzymes, such as quinoprotein methanol dehydrogenase in methylotrophs, are readily solubilized when the cell-free extract is prepared [2]. Given that oxidative fermentation is only functional under fairly acidic conditions, D -gluconate oxidation with an NADP-dependent enzyme observed at alkaline pH is unlikely to participate directly in keto- D -gluconate production under acidic conditions. Although FAD is linked covalently to FAD-dependent enzymes and PQQ is tightly bound to enzyme proteins (though all PQQ-dependent enzymes (quinoproteins) contain PQQ as dissociable form), most of the membrane-bound dehydrogenases indicated earlier were stable and active without exogenous addition of the responsible coenzyme, giving the impression that they were coenzymeindependent or NAD(P)-independent dehydrogenases. However, when the cellfree extract is centrifuged under stronger centrifugal force (e.g. 68 000 × g for 60 min), the enzyme activity of the membrane-bound enzymes precipitates as the membrane fraction, while the majority of NAD(P)-dependent enzymes exist in the supernatant. A typical membrane-bound dehydrogenase can be freed from NAD(P)-dependent enzyme activity by such simple fractionation of cell-free extract. The membrane-bound dehydrogenases are only solubilized by the aid of detergents. Addition of a chaotropic agent like KCl sometimes increases the recovery of solubilized enzyme. According to the physiological roles of membrane-bound and periplasmic enzymes, it should be beneficial to detoxify cellular toxic compounds such as methanol or amines outside cells (for example, in the periplasmic space) and not in the cytoplasm. When a cell-free extract is fractionated with ammonium sulfate, membrane-bound dehydrogenases tend to precipitate at relatively low concentrations of ammonium sulfate, in contrast to NAD(P)-dependent enzymes. Sometimes unexpectedly low enzyme recovery is seen after purification of cell-free extracts on ion exchange chromatographic column, suggesting the presence of membrane-bound enzymes.

1.2 Basic Technical Information Regarding Membrane-bound Enzymes 1.2.1 Preparation of Cytosolic Fractions and Membrane Fractions

A cell suspension is prepared by homogenizing freshly harvested cells at a ratio of about 10 g wet cells per 10 mL of appropriate buffer solution. For mechanical

1.2 Basic Technical Information Regarding Membrane-bound Enzymes

cell disruption, a cell suspension is passed through a French pressure cell press at 1000 kg cm−2 or other criteria. After removal of the intact cells by a conventional low-speed centrifuge, the crude cell-free extract is further centrifuged at 68 000 × g for 60 min to separate the membrane fraction as precipitate, and the resulting supernatant is designated as the soluble fraction. In the soluble fraction, it should be noted that enzyme activities of both cytoplasmic and periplasmic enzymes are usually observed. It is better to wash the resulting membrane precipitate by suspending and homogenizing in the same buffer by ultracentrifugation one or two times to eliminate the soluble enzymes. 1.2.2 EDTA Treatment of the Membrane Fraction Carrying PQQ as Coenzyme

After a membrane suspension (10 mg of protein per mL) is mixed with 20 mmol L −1 EDTA for 30 min in an ice bath. The excess EDTA is removed from the membrane by ultracentrifugation two or three times at 68 000 × g for 60 min. The precipitate is resuspended in a buffer to wash EDTA out of the membrane fraction, followed by ultracentrifugation again under the same conditions. The resulting precipitate is resuspended with the same buffer. Under these conditions, as mentioned below, many PQQ-dependent dehydrogenases are resolved to apoenzymes. However, some PQQ-dependent dehydrogenases are still active, although some decrease in enzyme activity is observed. If no loss of enzyme activity is found, the presence of covalently bound FAD as the coenzyme is the alternative possi-bility. To convert the apoenzyme to the holoenzyme, PQQ and divalent cations such as Ca2+ or Mg2+ are added to 5 μmol L −1 and 5 mmol L −1, respectively, and the enzyme incubated, for example, for 30 min at 25 ºC, until full enzyme activity returns. 1.2.3 Assays of Enzyme Activity

Most PQQ- and FAD-dependent dehydrogenases can be assayed using artificial electron acceptors such as potassium ferricyanide or phenazine methosulfate (PMS) [3]. In the case of potassium ferricyanide, enzyme activity can be assayed over a broad pH range, from highly acidic to highly alkaline conditions. On the other hand, enzyme activity measurement with PMS combined with dichlorophenol indophenol (DCIP) is invalid at acidic pH below 6 due to non-enzymatic decolorization of the electron acceptor used. Thus, the assay with PMS-DCIP is valid in the neutral to alkaline regions. It is also worth noting that when enzymes containing a heme c component in the enzyme molecule or membrane fraction are used, the enzyme activities can be easily assayed with potassium ferricyanide. However, enzyme activity measurement with PMS-DCIP is invalid if the enzymes do not contain the heme c component after solubilization from the membrane.

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1.3 PQQ-Dependent Dehydrogenases 1.3.1 Alcohol Oxidation 1.3.1.1

Membrane-Bound Alcohol Dehydrogenase (ADH III)

Ethanol → Acetaldehyde Quinohemoprotein alcohol dehydrogenase (ADH III) is localized on the outer surface of the cytoplasmic membrane of the acetic acid bacteria, Acetobacter and Gluconobacter. ADH III has the most important role in vinegar production in oxidizing ethanol to acetaldehyde. The acetaldehyde generated is immediately oxidized to acetate by aldehyde dehydrogenase located close to ADH III on the same cytoplasmic membrane. As mentioned elsewhere, most people still believe that ethanol oxidation during vinegar production must be catalyzed by a cytosolic NAD-dependent ADH, even though the discovery of ADH III was reported in 1960s. ADH III catalyzes alcohol oxidation under acidic conditions at pH 3–4, which is a favorable biological environment for vinegar production. It is distinct from the cytosolic NAD-dependent ADH that oxidizes alcohol under fairly alkaline conditions at pH 9–11. When acetic acid bacteria are growing on ethanol, the enzymic activity of the cytosolic NAD-dependent ADH is very weak and almost undetectable. Among a huge number of PQQ-dependent dehydrogenases, ADH III has been purified and demonstrated to be a typical example of a membrane-bound dehydrogenase [1]. It was the first membrane-bund enzyme to be crystallized (in 1982) [4]. The purified ADH III shows heme c type absorption spectra with absorption maxima at 555, 523, and 418 nm in the reduced enzyme. A shift in the γ-peak to 413 nm was observed for the oxidized enzyme. ADH III consists of three different subunits: subunit I (80 kDa), subunit II (50 kDa), and subunit III (20 kDa) (Fig. 1.3). Subunit I contains the catalytic site involving PQQ as the primary coenzyme and one heme c, forming a superbarrel structure surrounded by eight propeller structures based on tryptophan [5]. ADH III has been shown to donate electrons to ubiquinone embedded in the membrane phospholipids, and then to the terminal oxidase. Although no ADH III subunits have a transmembrane domain, subunit II (cytochrome subunit) has been shown to have a ubiquinone reacting site and to have two amphiphilic αhelices as a possible membrane anchor [6]. ADH III also seems to bind to the membrane and thus to transfer the reducing equivalent to ubiquinone via subunit II. Subunit II involves three heme c components and binds the ADH III to the cytoplasmic membrane [6, 7]. The physiological function of subunit III is still unknown but it is obviously important in the ADH III reaction. If subunit III is deleted, ethanol oxidation no longer takes place.

1.3 PQQ-Dependent Dehydrogenases

Fig. 1.3 Proposed structure of quinohemoprotein alcohol dehydrogenase (ADH III).

The ethanol oxidase system is composed of three simple components: ADH III, ubiquinone-9 (UQ9 ) in the case of Acetobacter or ubiquinone-10 (UQ10 ) in Gluconobacter, and the transmembrane terminal oxidase, which functions as an ubiquinol oxidase. The electron generated is transferred to UQ9 or UQ10 converting it to the reduced form, ubiquinol-9 or ubiquinol-10, which in turn are oxidized to UQ9 or UQ10 again. The terminal oxidase transfers the reducing equivalent across the cytoplasmic membrane, yielding a proton gradient (inside negative) allowing bioenergy generation. Thus, acetic acid bacteria are able to obtain bioenergy during substrate oxidation on the outer surface of the cytoplasmic membrane. This hypothesis has been confirmed using a reconstituted proteoliposome of the alcohol oxidase system involving ADH III, UQ9 or UQ10 and the terminal oxidase [8]. A clear proton gradient across an artificial proteoliposome membrane vesicle is formed by the addition of substrate and can be followed by quenching of fluorescent dye along with alcohol oxidation. For more details, refer to [1, 5]. ADH III catalyzes the irreversible oxidation of primary alcohols except for methanol. Alcohol oxidation is conducted with different types of catalyst containing ADH III such as growing cells (vinegar production), resting cells, isolated

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membrane fractions, and purified ADH III. In this context, ADH III has been use in the construction of alcohol sensors or alcohol biosensors that are able to interact directly with an electrode. Unlike the NAD(P)-dependent alcohol dehydrogenases, alcohol oxidation is catalyzed under fairly acidic conditions at pH 3–6. Ethanol-grown cells of Acetobacter or Gluconobacter show a strong ethanoloxidizing activity with the membrane fraction while a little enzyme activity of NAD(P)-dependent alcohol dehydrogenase is observed [9]. Ethanol oxidation with an NAD(P)-dependent alcohol dehydrogenase usually shows a pH optimum under highly alkaline conditions at pH 9–11 and aldehyde reduction to alcohol favorably occurs under acidic conditions at pH 5–6. On the other hand, if ADH III is deleted by means of genetic mutation, the enzyme activity of ADH III disappears in the membrane fraction and the cytosolic NAD(P)dependent ADH becomes predominant and the specific activity increased remarkably. Acetate production with such a mutant no longer takes place and the total cell growth increases, because ethanol added to the culture medium is fully converted to carbon and energy sources for the mutant [10]. Among strains of acetic acid bacteria, acetate overoxidation (also known as acetate peroxidation) is an unfavorable phenomenon in acetate brewery. Our understanding of the molecular mechanism and physiological aspects of acetate peroxidation have progressed [6, 11]. Structural studies of ADH III by means of X-ray crystallography are now being undertaken. In the case of ADH III of acetic acid bacteria, direct electron transfer to the electrode has been shown by direct binding of the enzyme to the hydrophobic surface of a gold electrode, which is able to transfer electrons directly to the electrode without any help of electron mediator [12, 13]. Such direct electron transfer to the electrode with ADH III has been confirmed in the case of the enzyme being entrapped onto a platinum electrode with polypyrrole [14]. This intermolecular electron transfer reaction has also been observed in two quinohemoproteins: ADH IIB (soluble quinohemoprotein ADH, see below) and ADH III. Based on the crystal structure of ADH IIB [15–17], it can be seen that the heme c domain in the C-terminus of the primary structure is distant from the Nterminal PQQ domain, which is highly homologous to the superbarrel structure of other PQQ-dependent enzymes [5], and covers the PQQ domain. The distance between the heme c and PQQ is about 20 Å in the reduced form, which seems to be a little far for direct electron transfer from PQQ to heme c. Therefore, the electron transfer from PQQ to heme c could occur via an amino acid backbone present between both domains, or directly, upon a conformational change due to oxidation. The same kind of electron transfer may be expected in the quinohemoprotein subunit of ADH III. Furthermore, in the multimeric enzyme it has been suggested that electrons extracted from ethanol at the PQQ site could be transferred via the heme c site in the subunit I to one of three heme c sites in the cytochrome subunit (subunit II), and the electrons passed to ubiquinone through two of the heme c moieties (Fig. 1.3) [18, 19]. In the case of ADH III, subunit II has been shown to have a ubiquinone reacting site despite having no transmembrane domain. Thus, ubi-


quinone reduction in ADH III may occur via electron transfer from PQQ through three of the four heme c moieties present. The electron transfer kinetics of quinoproteins or quinohemoproteins may become critical from the biotechnological point of view, especially when applying them to alcohol biosensors.

1.3.1.2

Soluble Alcohol Dehydrogenases

Alcohol → Aldehyde A bacterial strain that is able to utilize several kinds of alcohols as its sole carbon and energy sources was isolated from soil and tentatively identified as Pseudomonas putida HK5 [20]. Three distinct dye-linked ADHs, each of which contains PQQ as the prosthetic group, are formed in the periplasmic space of the strain grown on different alcohols and thus the enzyme activities appear in the cell-free extract in soluble form. ADH I is formed most abundantly in the cells grown on ethanol and is similar to the quinoprotein ADH reported from P. putida [21], except for its isoelectric point. The other two ADHs, ADH IIB and ADH IIG, are formed separately in the cells grown on 1-butanol and 1,2-propandiol, respectively. Both of these enzymes contain heme c in addition to PQQ and function as quinohemoprotein dehydrogenases. Thus, potassium ferricyanide is an available electron acceptor for ADHs IIB and IIG but not for ADH I. The molecular masses are estimated to be 69 kDa for ADH IIB and 72 kDa for ADH IIG, and both enzymes have been shown to be monomers. Antibodies raised against each of the purified ADHs could distinguish the ADHs from one another. Immunoblot analysis showed that ADH I is detected in cells grown on each alcohol tested, but ethanol was the most effective inducer. ADH IIB is formed in cells grown on alcohols of medium chain length and also on 1,3-butanediol. Induction of ADH IIG is restricted to 1,2-propanediol or glycerol, of which the former alcohol is more effective. These results from immunoblot analysis correlate well with the substrate specificities of the respective enzymes. Thus, three distinct quinoprotein ADHs are formed by a single bacterium under different growth conditions. Molecular cloning and structural analysis of ADH IIB and ADH IIG have been reported [16, 17, 22]. These soluble forms of ADHs are also used for alcohol sensors. Of the three different PQQ-dependent ADHs, ADH IIG would be the most convincing for use as a glycerol sensor for a neutral fat diagnostic test by combining with lipoprotein lipase.

1.3.1.3 Cyclic Alcohol Dehydrogenase (Secondary Alcohol Dehydrogenase), Membrane-Bound

Cyclic alcohol → Cyclic ketone Aliphatic secondary alcohol → Aliphatic ketone

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Numerous investigations on microbial and enzymatic oxidation of cyclic alcohols have been carried out with various bacteria. Most of them are related to NADdependent cyclohexanol dehydrogenase (EC 1.1.1.245) from pseudomonads and Acinetobacter sp. [23–26]. Microbial cyclopentanol oxidation was also studied in Pseudomonas sp. and NAD-dependent cyclopentanol dehydrogenase (EC 1.1.1.163) was indicated as the responsible enzyme in the first step of the oxidative metabolism of cyclic alcohol [27]. Cyclohexanol oxidation was also observed by a secondary alcohol oxidase from Pseudomonas sp. in which hydrogen peroxide-producing flavin-dependent enzyme is functional [28–30]. The microorganisms described in the abovementioned papers do not accumulate oxidation products outside the cells, because most enzymes dealt with are cytoplasmic enzymes. Nothing has been reported, therefore, about membrane-bound cyclic alcohol dehydrogenase located on the outer surface of the cytoplasmic membrane allowing accumulation of the oxidation products outside the cells. A quinoprotein catalyzing oxidation of cyclic alcohols was found in the membrane fraction of acetic acid bacteria. After extensive screening, Gluconobacter frateurii CHM 9 was selected. EDTA treatment with the membrane fraction indicated that the membrane-bound cyclic alcohol dehydrogenase is a PQQdependent dehydrogenase. From the membrane fraction, PQQ-dependent cyclic alcohol dehydrogenase was purified [31]. In contrast, from the cytoplasmic fraction of the same organism, an NAD-dependent cyclic alcohol dehydrogenase was purified and crystallized. The substrate specificities of the two differently localized enzymes showed an interesting contrast with each other, suggestive of their different physiological roles in the organisms. Unlike the already known cytosolic NAD(P)H-dependent alcohol-aldehyde or alcohol-ketone oxidoreductases, the PQQ-dependent enzyme is unable to catalyze reduction of cyclic ketones or aliphatic ketones to cyclic alcohols or aliphatic secondary alcohols. PQQ-dependent cyclic alcohol dehydrogenase was solubilized from the membrane fraction by differential solubilization to eliminate the predominantly existing ADH III and the two subsequent steps of column chromatography gave a homogeneous enzyme preparation. The purified PQQ-dependent enzyme has a molecular mass of 83 kDa in SDS-PAGE. A wide variety of cyclic alcohols are oxidized with the PQQ-dependent enzyme. Once the enzyme was solubilized, only the PMS-DCIP assay was valid, because the enzyme does not contain any heme c component. When the PQQ-dependent enzyme was compared with the cytosolic NAD-dependent enzyme, the reaction rate and the total enzyme activity found in cyclic alcohol oxidation with the PQQ-dependent enzyme was more than 100 times higher than that with the cytosolic NAD-dependent enzyme. The NAD-dependent enzyme is unfavorable for oxidation reactions, but is favorable for the reduction of cyclic ketones to cyclic alcohols or aliphatic ketones to aliphatic secondary alcohols. Thus, the NAD-dependent enzyme makes no contribution to cyclic alcohol oxidation, and reduction of cyclic ketones and aliphatic ketones to their corresponding alcohols is favored in the presence of NADH.


1.3.2 Glucose Oxidation 1.3.2.1

Membrane-Bound D -Glucose Dehydrogenase (m-GDH) D -Glucose

→ D -glucono-δ-lactone

The quinoprotein D -glucose dehydrogenase (EC 1.1.99.17) (GDH) occurs on the outer surface of the cytoplasmic membrane of oxidative bacteria such as Pseudomonas and Gluconobacter strains and catalyzes direct oxidation of D -glucose to D gluconate via D -glucono-δ-lactone (membrane-bound D -glucose dehydrogenase, m-GDH). It is known as an alternative pathway to the phosphotransferase system of bacteria to catalyze D -glucose assimilation. m-GDH is found in a variety of bacteria including Gram-negative facultative anaerobes such as enteric bacteria and Zymomonas, as well as aerobic bacteria such as pseudomonads and acetic acid bacteria. GDH was originatally investigated in Acinetobacter calcoaceticus in the early 1960s by Hauge [32–35] and subsequently in the late 1970s by Duine et al. [36], who showed the enzyme to be a quinoprotein. A. calcoaceticus contains a soluble form of GDH (s-GDH) in addition to a membrane-bound form (m-GDH). Thus, they had been believed for many years to be the same enzyme or interconvertible forms. However, recent evidence has shown that this soluble enzyme is not a typical m-GDH. The membranes from several strains such as Escherichia coli, Klebsiella aerogenes, Pseudomonas aeruginosa, Gluconobacter suboxydans, Acetobacter aceti, and A. calcoaceticus contain antigens cross-reactive with an antibody of m-GDH purified from P. fluorescens [37], while s-GDH purified from the soluble fraction of A. calcoaceticus does not cross-react with the antibody [38]. Subsequently, s-GDH and m-GDH were purified separately from A. calcoaceticus and shown to be distinctive in all aspects, including optimum pH, kinetics, substrate specificity, ubiquinone reactivity, molecular size, and immunoreactivity [39]. As described below, s-GDH of A. calcoaceticus is a monomer consisting of a single polypeptide of 48–55 kDa containing one molecule of PQQ. When m-GDH was solubilized and purified from Gluconobacter suboxydans IFO 12528, other species of membrane-bound dehydrogenases were eliminated by treating the enzyme solution at pH 2.5 in the initial stage of enzyme purification. The purified m-GDH was homogeneous in analytical ultracentrifugation (4.2 S) and sucrose density gradient centrifugation [40]. m-GDH from acetic acid bacteria was highly hydrophobic and 87 kDa of its molecular mass has been determined by SDS-PAGE in the presence of urea. The existence of PQQ as the primary coenzyme has been confirmed with the purified enzyme. The optimum pH of D -glucose oxidation is found to be pH 3.0 with potassium ferricyanide and pH 6.0 with PMS-DCIP. The substrate specificity of the enzyme seems to be restricted to D -glucose, and other sugars are not oxidized except for maltose, which is oxidized at a low rate. Due to the hydrophobicity, the enzyme is regarded as a typical integral membrane protein in acetic acid bacteria.

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Since E. coli does not have the PQQ gene, the bacterium produces m-GDH in the apo-form. The apoenzyme was converted to active holoenzyme by the addition of PQQ and Mg2+ [41]. In contrast to the m-GDH found in acetic acid bacteria [40], PMS-DCIP is a convenient electron acceptor for measuring D -glucose oxidation by this enzyme. The enzyme is a typical membrane-bound enzyme highly embedded in the cytoplasmic membrane, with five membrane-spanning domains [42. 43]. The active site of m-GDH faces the periplasmic space. m-GDH was solubilized from the cytoplasmic membrane with the aid of detergent and further purified to a homogeneous state. The substrate specificity of E. coli m-GDH looks similar to that of A. calcoaceticus [39]. D -Gluconate is an important substance for the food industry as a food additive. Ca-D -gluconate is widely used as a metal polish in the lens industry. In aerobic Gram-negative bacteria including acetic acid bacteria, D -gluconate is produced from D -glucose by the action of m-GDH. It is distinct from a fungal D -glucose oxidase producing D -gluconate that the bacterial enzyme does not link to molecular oxygen directly while the fungal enzyme utilizes molecular oxygen-producing hydrogen peroxide. m-GDH has been shown to donate electrons to ubiquinone, embedded in the membrane phospholipids, and then to the terminal oxidase [1]. Unlike the previous assumption, however, m-GDH does not seem to have a ubiquinone reacting site in the deeply embedded region of the membrane. Despite having a transmembrane domain, m-GDH has been shown to react with ubiquinone near the surface of the membrane [44]. This notion has been strengthened by the recent finding that the C-terminal half of E. coli m-GDH, the N-terminal transmembrane domain being deleted, maintains ubiquinone reductase activity [45]. To clarify the intermolecular electron transfer of m-GDH, quantitation and identification of ubiquinone have been done, indicating that E. coli m-GDH contains a tightly bound ubiquinone-8 (UQ8 ) in its molecule. A significant increase in the EPR signal was observed following D -glucose addition in m-GDH reconstituted with PQQ and Mg2+ after the addition of D -glucose, suggesting that bound UQ8 accepts a single electron from PQQH2 to generate semiquinone radicals. No such increase in the EPR signal was observed in UQ8 -free m-GDH under the same conditions. Moreover, a UQ2 reductase assay with a ubiquinone-related inhibitor (C49) revealed different inhibition kinetics between the wild-type m-GDH and UQ8 -free m-GDH. It is proposed that the native m-GDH bears two ubiquinonebinding sites, one (QI) for bound UQ8 in its molecule and the other (QII) for UQ8 in the ubiquinone pool, and the bound UQ8 in the QI site acts as a single electron mediator in the intramolecular electron transfer in m-GDH [46]. It should be noted that there is no specific D -xylose dehydrogenase other than m-GDH. D -Xylose oxidation is done by m-GDH. 1.3.2.2

Soluble D -Glucose Dehydrogenase (s-GDH) D -Glucose

→ D -glucono-δ-lactone

As mentioned above, A. calcoaceticus contains s-GDH in addition to m-GDH. s-GDH purified from the soluble fraction of A. calcoaceticus does not cross-react


with the antibody for m-GDH [38]. Subsequently, s-GDH and m-GDH were purified separately from A. calcoaceticus and shown to be distinctive in all aspects including optimum pH, kinetics, substrate specificity, ubiquinone reactivity, molecular size, and immunoreactivity [39]. s-GDH of A. calcoaceticus is a monomer consisting of a single polypeptide of 48–55 kDa containing one PQQ molecule. A gene (gdhB) for the enzyme has been cloned and the gene product is estimated to be a polypeptide of 52.8 kDa that contains a 24-amino-acid signal sequence at its N-terminus and thus it is expected to become the mature protein of 50.2 kDa having no hydrophobic regions. Thus, s-GDH seems to be translocated through the cytoplasmic membrane into the periplasmic space. s-GDH is capable of catalyzing the oxidation of disaccharides, lactose, or maltose, as well as D -glucose. The enzyme is able to donate electrons to several artificial dyes, including PMS, DCIP as well as short-chain ubiquinone homologs, Q1 and Q2, but it is unable to react with the longer chain ubiquinones Q6 and Q9 [39]. s-GDH was purified from the soluble fraction of A. calcoaceticus with the overall recovery of 28% and showed a final specific activity of 2210 units mg−1. The substrate specificity of s-GDH is different from that of m-GDH. s-GDH oxidizes D -glucose (100%), D -fucose (28%), D -xylose (15%), D -galactose (30%), maltose (93%), and lactose (72%), while m-GDH is less reactive with disaccharides: D -glucose (100%), D -fucose (119%), D -xylose (81%), D -galactose (73%), D -ribose (54%), maltose (13%), and lactose (5%). 1.3.2.3 Applications of Quinoprotein GDHs as D -Glucose Sensors As our understanding of the mechanisms of catalysis and electron transfer of quinoproteins progresses, the idea of the application of these enzymes as biosensors becomes more attractive, and thus electrochemical studies aimed at the design of specific quinoprotein-based electrodes have increased. s-GDH is the first quinoprotein applied to such a biosensor, and a D -glucose sensor with s-GDH in a single-use electrochemical test strip containing ferricyanide as a mediator is already on the market. Prior to the appearance of this s-GDH sensor, a fungal flavoprotein D -glucose oxidase and NAD-dependent GDH had been used as the enzyme for D -glucose biosensors. However, biosensors based on D -glucose oxidase are sensitive to oxygen fluctuations in blood samples, while those based on NAD GDH have poor stability due to the loss of the cofactor NAD. These problems are overcome by using PQQ-dependent GDH, which is oxygen insensitive and has a tightly bound PQQ. In addition, since the electron transfer rate of s-GDH is very much higher than that of D -glucose oxidase, the biosensor based on s-GDH has produced more than twice the current density of similar D -glucose oxidase-based electrodes when s-GDH is immobilized in the presence of an insoluble ferrocene [47]. Thus, several characteristics of the s-GDH-based D -glucose sensor, especially the high catalytic activity and oxygen insensitivity, make it suitable for in vivo blood D -glucose monitoring in the management of diabetes. Although it has still not been applied to commercial sensors, m-GDH has been examined for its electrochemical reaction on electrodes. Attempts have been

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made to attach m-GDH from Erwinia sp. to an electrode surface using several different materials such as carbon paste, where no direct electron transfer from the enzyme is observed unless some insoluble quinone mediator is included in the paste [48]. Anodic current has been detected with Erwinia m-GDH depending on D -glucose concentrations, and is greatly increased in the presence of soluble electron mediators such as PMS. In order to improve the specificity and the response range toward D -glucose, the same type of m-GDH-based electrode has been prepared by immobilizing m-GDH from an E. coli mutant, in which His775 is substituted with Asp, on the carbon paste electrode [49]. The mutated m-GDHentrapped electrode exhibits an expanded response range for D -glucose (3– 70 mmol L −1) and a narrower substrate specificity. The substrate specificity of m-GDH from G. suboxydans IFO 12528 is attractive for D -glucose biosensors. The enzyme is highly specific to D -glucose and maltose is oxidized by only 5% of that for D -glucose. Other aldohexoses and aldopentoses have no effects on m-GDH from acetic acid bacteria [40]. 1.3.3 Polyol Oxidation 1.3.3.1

D -Arabitol

D -Arabitol

Dehydrogenase, Membrane-Bound

→ D -xylulose

Because there is little information about C-5 sugar alcohol oxidation, evidence has been presented confirming that L-ribulose formation, the oxidation product of ribitol, was catalyzed by the action of a membrane-bound PQQ-dependent ribitol dehydrogenase, but not by a cytosolic NAD-dependent ribitol dehydrogenase [50]. Due to the high hydrophobicity and instability of the enzyme for pentitol oxidation, its solubilization from the membrane and purification remained to be achieved. Several physiological and catalytic properties of the purified membrane-bound D -arabitol dehydrogenase (ARDH) have been examined [51]. Solubilization of ARDH from the membrane of Gluconobacter suboxydans IFO 3257 was done successfully with Mydol 10. Selection of a favorable detergent, keeping ARDH as the holoenzyme during all the purification steps by the presence of PQQ and Ca2+ , and using a buffer system involving acetate buffer supplemented with Ca2+ were necessary to treat the highly hydrophobic and thus labile enzyme. Purification of ARDH was successful after two steps of column chromatography on DEAEToyopearl and CM-Toyopearl in the presence of detergent and Ca2+ . The purified ARDH was homogeneous and showed a single sedimentation peak in analytical ultracentrifugation at 3.6 S of the apparent sedimentation constant. Upon SDS-PAGE, ARDH dissociated into two different subunits of 82 kDa (subunit I) and 14 kDa (subunit II), forming a heterodimeric structure. It contained no heme component, unlike ADH IIB, ADH IIG, and ADH III. ARDH did not react with primary alcohols. The enzyme was proved to be a quinoprotein and


dissociation of PQQ was detected by HPLC by SDS-treated ARDH. PQQ and ubiquinone-10 (UQ10 ) were detected in a purified ARDH when enzyme solubilization was done with dodecyl-β-maltoside instead of Triton X-100. More importantly, when the membrane fraction was treated with 20 mmol L −1 EDTA overnight, ARDH activity was lost but the enzyme activity was restored to its original level by the subsequent addition of PQQ and Ca2+ . ARDH from G. suboxydans IFO 3257 was found to be a versatile enzyme for the oxidation of various sugar alcohols to the corresponding oxidation products, such as glycerol to dihydroxyacetone, ribitol to L-ribulose, D -arabitol to D -xylulose, D -sorbitol to L-sorbose, D -mannitol to D -fructose, and, surprisingly, D -gluconate to 5-keto- D -gluconate, according to the Bertrand–Hudson’s rule [52] (Fig. 1.4). Membrane-bound glycerol dehydrogenase (GLDH) from Gluconobacter industrius IFO 3260 was reported to have the same wide substrate specificity as ARDH [53]. However, due to the hydrophobicity, GLDH was unstable when purified and thus insufficient data were obtained to compare it with other PQQ-dependent dehydrogenases, although GLDH has been proved to be an enzyme containing PQQ as the coenzyme. ARDH is very similar to GLDH as well as the membranebound quinoprotein D -sorbitol dehydrogenase (SLDH) from G. suboxydans IFO 3255 [54] in the following respects: broad substrate specificity to sugar alcohols, absence of heme component in the enzyme, and a molecular mass of 75–80 kDa in SDS-PAGE. It is worth noting that the enzyme catalyzes D -gluconate oxidation to yield 5-keto- D -gluconate, whereas 2-keto- D -gluconate is produced by an FADdependent D -gluconate dehydrogenase (GADH). The oxidation product of D -gluconate with ARDH was identified as 5-keto- D gluconate, when purified ARDH was used for cyclic oxidation of D -gluconate in the presence of quinol oxidase and ubiquinone Q2 as illustrated in Fig. 1.5. The

Fig. 1.4 Bertrand–Hudson’s rule in polyol oxidation.

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Fig. 1.5 D -Gluconate oxidation by a coupling reaction consisting of polyol dehydrogenase and quinol oxidase.

reaction mixture was only reactive with 5-keto- D -gluconate reductase [55] but not with 2-keto- D -gluconate reductase [56], and a stoichiometric amount of 5-keto-D gluconate was detected in the reaction mixture. Thus, it was concluded that oxidative fermentation of 5-keto- D -gluconate is catalyzed by polyalcohol dehydrogenase and there is no specific 5-keto- D -gluconate-yielding D -gluconate dehydrogenase. Of course, in contrast to what had previously been believed, a cytosolic 5-keto- D gluconate reductase cannot possibly contribute to the oxidative fermentation producing 5-keto- D -gluconate. Regarding the enzyme structure, ARDH may be composed of two different subunits. The presence of a small open reading frame, which is very similar to the transmembrane region of m-GDH in E. coli, has been shown to be essential for the expression of active SLDH (Hoshino et al., personal communication). Therefore, as in SLDH, subunit II may be buried in the cytoplasmic membrane anchoring ARDH to the outer surface of the cytoplasmic membrane. Because the purified ARDH had a relatively high Q2 reductase activity of 12.3 units per mg of protein, ARDH seemed to be attached to the cytoplasmic membranes in vivo and to link to their electron transfer chain via ubiquinone (Adachi et al., unpublished data). In acetic acid bacteria, sugar alcohol oxidation by the membranebound enzyme is in accordance with Betrand–Hudson’s rule (Fig. 1.4), that is to say, according to the generalization by Bertrand, the most favorable confi guration for oxidation has the erythro form and R-configuration of two secondary hydroxyl groups adjacent to the primary alcohol group. ARDH oxidized some secondary alcohols, which must have an R-configuration. 1.3.3.2

meso-Erythritol Oxidation Dehydrogenase, Membrane-Bound

meso-Erythritol → L-erythrulose There is some information about the oxidative fermentation of C4 sugar alcohols in the literature, except earlier reports about meso-erythritol oxidation by acetic acid bacteria [57, 58]. Because L-erythrulose is not readily available from commercial sources, it is important to investigate the fermentation profi le of Lerythrulose production for the identification of the enzyme responsible for meso-erythritol oxidation, and to purify and characterize the enzyme to provide basic information on L-erythrulose production. According to the recommendation of the US Food and Drug Administration (FDA), dihydroxyacetone should be


replaced by L-erythrulose in cosmetics for those who are sensitive to dihydroxyacetone. Gluconobacter frateurii CHM 43 has been screened and shows high Lerythrulose production from meso-erythritol. NAD(P)-independent enzymes catalyzing meso-erythritol oxidation from the membrane fraction and NAD(P)dependent enzymes from the cytosolic fraction have been purified from the organism [59]. The purified enzyme from the membrane fraction was identified as a quinoprotein and is responsible for L-erythrulose production, but the NAD(P)dependent enzyme was independent for the production of L-erythrulose. Growing cells and the membrane fraction of the strain rapidly oxidized meso-erythritol to L-erythrulose irreversibly with almost 100% recovery at 37 ºC. L-Erythrulose was also produced efficiently by the resting cells as well. The enzyme responsible for meso-erythritol oxidation was localized on the outer surface of the cytoplasmic membrane of the organism. The prosthetic group of the enzyme was identified to be PQQ after inactivation of the enzyme by EDTA treatment and the subsequent restoration to original levels by the exogenous addition of PQQ or PQQ and CaCl2. The enzyme was solubilized and purified to homogeneity [59]. The purified enzyme showed a single band in SDS-PAGE with a molecular mass corresponding to 80 kDa. A smaller band at 10–20 kDa that anchors the enzyme to the cytoplasmic membrane was not detected. The optimum pH of meso-erythritol oxidation was found to be pH 5.0. The Michaelis constant of the enzyme for meso-erythritol oxidation was found to be 25 mmol L −1. The enzyme showed broad substrate specificity toward C3 –C6 sugar alcohols in which the erythro form of two hydroxy groups existed adjacent to the primary alcohol group according to the Bertrand–Hudson rule (Fig. 1.4). NAD(P)-dependent meso-erythritol dehydrogenase was purified to a crystalline state. The molecular mass was estimated to be 60 kDa, composed of two identical subunits. Unlike the membrane-bound quinoprotein, the enzyme catalyzes reversible oxidoreduction at an optimum pH of 9.0–10.5 for meso-erythritol oxidation and pH 6.0 for L-erythrulose reduction. It is evident that NAD(P)-dependent enzymes have no function in L-erythrulose production. 1.3.3.3

D -Gluconate

D -Gluconate

Oxidizing Polyol Dehydrogenase, Membrane-Bound

→ 5-keto- D -gluconate

Gluconobacter species oxidize sugars and sugar acids and uniquely accumulate two different keto- D -gluconates, 2-keto- D -gluconate, and 5-keto- D -gluconate, in the culture medium by the oxidation of D -gluconate [51, 60, 61]. Recently, PQQdependent ARDH and SLDH have been purified from G. suboxydans, both of which have similar broad substrate specificity towards several different polyols. ARDH and SLDH were shown to be identical based on their immunocrossreactivity and also on gene disruption and were suggested to be the same as the previously isolated glycerol dehydrogenase (GLDH) (EC 1.1.99.22). Thus, GLDH

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is the major polyol dehydrogenase involved in the oxidation of almost all sugar alcohols in Gluconobacter sp. In addition, the so-called quinoprotein GLDH was also uniquely shown to oxidize D -gluconate, which was completely different from flavoprotein D -gluconate dehydrogenase (GADH) (EC 1.1.99.3), that is the direct catalyst for the production of 2-keto- D -gluconate. During the investigation of ARDH, it was found that D arabitol oxidation was always parallel to D -gluconate oxidation. The gene disruption experiment and the reconstitution of the purified enzyme clearly showed that the production of 5-keto- D -gluconate in G. suboxydans is solely dependent on the quinoprotein GLDH (Fig. 1.6). Production of 5-keto- D -gluconate is important, allowing the Gray’s method for vitamin C production via L-idonate and 2-keto-L-gulonate to be practical. This pathway looks stable unlike the other well-known routes involving L-sorbosone or 2,5-diketo- D -gluconate as the intermediates. 5-Keto- D -gluconate is a stable compound while L-sorbosone and 2,5-diketo- D -gluconate are labile compounds (Fig. 1.7). Exclusive 5-keto-D -gluconate production would be possible using a mutant in which GADH yielding 2-keto-D -gluconate is deleted. As new information on 5-ketoD -gluconate production accumulates, 5-keto- D -gluconate production should become practical, which will allow us to utilize the new route for vitamin C production proposed by Gray [62, 63]. According to this, half of the 5-keto- D -gluconate is converted to L-idonate and then to 2-keto-L-gulonate by another oxidative fermentation before finally being converted to L-ascorbate, as shown in Fig. 1.7.

Fig. 1.6 Comparison of keto- D -gluconate accumulation with a wild strain (left frame) and a mutant lacking polyol dehydrogenase (right frame). G. suboxydans IFO 3255 wildtype strain (left frame) and GLDH-defective mutant 3255sldA::Km (right frame) were cultured in 100 mL of D -glucose– D -gluconate medium. During cultivation, 2KGA and 5KGA were analyzed periodically as indicated.


Fig. 1.7 Different routes for vitamin C.

1.3.3.4

Glycerol Dehydrogenase, Membrane-Bound

Glycerol → Dihydroxyacetone Glycerol oxidation to dihydroxyacetone in acetic acid bacteria is catalyzed by the membrane-bound quinoprotein glycerol dehydrogenase (GLDH), synonymous to polyol dehydrogenase, which catalyzes direct oxidation of various kinds of polyols according to the Bertrand–Hudson rule (Fig. 1.4). Since Bertrand reported the oxidation of glycerol to dihydroxyacetone by a sorbose bacterium that he had isolated, the production of dihydroxyacetone by oxidative fermentation advanced [60], although the nature of the responsible enzyme for dihydroxyacetone production remained unclear. It was in 1985 that the first investigation of the membrane-bound GLDH of Gluconobacter industrius IFO 3260 was carried out [53]. In most acetic acid bacteria, the enzyme activity of ADH III is predominant in the cytoplasmic membrane and GLDH is readily solubilized together with ADH III. Once GLDH is solubilized, it is quite difficult to separate GLDH from ADH III under common techniques of chromatography. Thus, for the solubilization and purification of GLDH, G. industrius IFO 3260 was selected, although the total enzyme activity of GLDH was not as high the strain showed the least enzyme activity of ADH III.

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For dihydroxyacetone production by growing cells, strains such as G. cerinus IFO 3264, IFO 3268, G. gluconicus IFO 3171, G. suboxydans IFO 3286, G. suboxydans IFO 3291, and G. suboxydans IFO 12528 would give better yields of dihydroxyacetone from glycerol. Since GLDH is highly hydrophobic and seems to be deeply embedded in the cytoplasmic membrane, the improved stability of GLDH after solubilization with detergents became the immediate subject to which a clear solution was required. Among various detergents tested, 0.5% of dimethyldodecylamineoxide was chosen as a favorable detergent for GLDH solubilization and GLDH purification was done simply by fractionation with polyethyleneglycol 6000 (PEG). The precipitate obtained with PEG from 15 to 25% gave 100 times increase in the specific activity from the membrane fraction, giving a final yield of 25%. The purified enzyme was transparent and had no characteristic color, unlike ADH III. During enzyme purification, contamination by other species of membranebound dehydrogenases found in the same membrane fraction of G. industrius, such as alcohol, D -gluconate, D -fructose, and aldehyde dehydrogenases, was completely eliminated. This was checked by substrate specificity. The oxidation rate of glycerol compared with that of other polyols such as meso-erithritol, D -arabitol, or D -sorbitol settled down to a constant level throughout the purification, while the enzyme activity toward D -glucose, D -fructose, or D -gluconate disappeared. When GLDH purification was tried, the numbers of detergents available for enzyme solubilization were restricted. If GLDH purification could be done with the other recently developed detergents such as octyl-β-glucoside, dodecyl-βmaltoside or a series of Mydol, stability would be much improved as is seen with D -arabitol dehydrogenase (ARDH) or meso-erythritol dehydrogenase. From the results obtained with GLDH, it is highly probable that oxidation of different polyols is catalyzed in acetic acid bacteria with an enzyme like GLDH. GLDH may be used for a diagnostic sensor monitoring glycerol generated from neutral fat after hydrolysis with lipoprotein lipase. Since GLDH catalyzes irreversible oxidation of glycerol to dihydroxyacetone and there is no reaction equilibrium, even a trace amount of glycerol generated can be measured with high accuracy, unlike other NAD(P)-dependent enzymes. 1.3.3.5

D -Mannitol

D -Mannitol


→ D -fructose

D -Mannitol oxidation to D -fructose in acetic acid bacteria is catalyzed by the membrane-bound PQQ-dependent polyol dehydrogenase, similar to GLDH or ARDH. The oxidation of D -mannitol to D -fructose was reported in many ketogenic acetic acid bacteria, as in the case of D -sorbitol oxidation. Fulmer and Underkofler looked at the possibility of using this oxidation for the industrial producton of D fructose using A. suboxydans (G. suboxydans at present) [64]. More detailed study of polyol dehydrogenase from acetic acid bacteria was carried out by Arcus and


Edson [65]. Two different enzymes were indicated in the organism. One enzyme was an NAD(P)-independent particulate dehydrogenase active on D -mannitol and other polyols at pH 5.0, with a substrate specificity following Bertrand–Hudson’s rule (Fig. 1.4). The other enzyme was NAD(P)-dependent. As shown in other cases in this chapter, NAD(P)-dependent dehydrogenases have no use for oxidative fermentation. Thus, only the membrane-bound enzyme contributes to D mannitol oxidation to D -fructose. In our recent study, several strains of thermotolerant acetic acid bacteria that can grow at 37–40 ºC were screened. Typical mesophilic strains cannot grow at these temperatures. Among the thermotolerant strains, Gluconobacter frateurii CHM 16 was chosen as the best biocatalyst for D -fructose production with the highest conversion efficiency as well as the highest yield [66]. Purification of the enzyme catalyzing D -mannitol oxidation was not tried, due to the issues described above. If we tried this, we would get a polyol dehydrogenase showing broad substrate specificity as seen with other examples in this chapter. 1.3.3.6

Ribitol Dehydrogenase, Membrane-Bound

Ribitol → L-ribulose Ribitol oxidation to L-ribulose in acetic acid bacteria is catalyzed by the membrane-bound PQQ-dependent polyol dehydrogenase, similar to GLDH or ARDH. Membrane-bound ribitol dehydrogenase in acetic acid bacteria catalyzes L-ribulose formation, while an NAD-dependent ribitol dehydrogenase is independent of oxidative fermentation [50]. To identify the enzyme responsible for pentitol oxidation by acetic acid bacteria, two different ribitol-oxidizing enzymes, one NAD(P)-dependent in the cytosolic fraction and the other NAD(P)-independent in the membrane fraction, were examined with respect to the oxidative fermentation. The cytoplasmic NAD-dependent ribitol dehydrogenase (EC 1.1.1.56) was crystallized from Gluconobacter suboxydans IFO 12528 and found to be an enzyme of molecular mass 100 kDa and 5 S as the sedimentation constant, composed of four identical subunits of 25 kDa each. The enzyme catalyzed a shuttle reversible oxidoreduction between ribitol and D -ribulose in the presence of NAD and NADH, respectively. Xylitol and L-arabitol were well oxidized by the enzyme with reaction rates comparable to ribitol oxidation. D -Ribulose, L-ribulose, and L-xylulose were well reduced by the enzyme in the presence of NADH as cosubstrates. The optimum pH of pentitol oxidation was found to be at alkaline pH such as 9.5–10.5 and ketopentose reduction was optimum at pH 6.0. NAD-Dependent ribitol dehydrogenase seemed to be specific to oxidoreduction between pentitols and ketopentoses. D -Sorbitol and D -mannitol were not oxidized with the NAD-dependent enzyme. However, no D -ribulose accumulation was observed outside the cells during the growth of the organism on ribitol. LRibulose was only accumulated in the culture medium instead, as the direct oxidation product catalyzed by the membrane-bound NAD(P)-independent ribitol

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dehydrogenase. Thus, the physiological role of NAD-dependent ribitol dehydrogenase seems to be to catalyze ribitol oxidation to D -ribulose in the cytoplasm. Phosphorylated D -ribulose is involved in the pentose phosphate pathway. LRibulose outside the cells could be incorporated into the cytoplasm in several ways when the use of L-ribulose as carbon and energy source becomes necessary for cell survival. From a series of simple experiment, membrane-bound PQQdependent sugar alcohol dehydrogenase was concluded to be the enzyme responsible for L-ribulose production from ribitol in oxidative fermentation by acetic acid bacteria. 1.3.3.7

D -Sorbitol

D -Sorbitol


→ L-sorbose

D -Sorbitol oxidation to L-sorbose in acetic acid bacteria is catalyzed by the membrane-bound PQQ-dependent polyol dehydrogenase in addition to FADdependent D -sorbitol dehydrogenase (SLDH) (see Section 1.4.5). The introduction of D -sorbitol oxidation to L-sorbose by microbial bioconversion (L-sorbose fermentation) stimulated the vitamin C industry [60]. The chemical method for L-ascorbate synthesis developed by Leichstein in 1937 leads to racemic sorbose after oxidation of D -sorbitol, while L-sorbose fermentation gives only L-sorbose with almost 100% yield, apparently two times higher than the chemical method. Many strains of “sorbose bacterium” have been isolated and used for L-sorbose production. Together with acetate production this process is a typical and classical microbial bioconversion with acetic acid bacteria. However, the enzymatic mechanism of L-sorbose formation remained to be clarified. In 1982, SLDH was purified for the first time from the membrane fraction of Gluconobacter suboxydans var. α IFO 3254 [67]. The enzyme contains a covalently bound FAD as the coenzyme, as described below (see Section 1.4.5). Until recently, before we started to survey the enzymatic properties of polyol dehydrogenases involved in D -sorbitol metabolism in acetic acid bacteria, SLDH from G. suboxydans var. α IFO 3254 was the sole described enzyme [50, 68, 69]. Through investigations on quinoproteins such as GLDH, ARDH, meso-erythritol dehydrogenase, and ribitol dehydrogenase, it has been concluded that these enzymes are responsible for D -sorbitol oxidation to L-sorbose as described in this chapter. Recently, we have screened thermotolerant acetic acid bacteria that can grow at 37 ºC. Gluconobacter frateurii CHM 54 was isolated and applied to L-sorbose production [66]. In most cases, strict temperature control is required for oxidative fermentation. A hot summer readily allows indoor temperatures to rise above 30 ºC in many countries. This is a serious challenge not only to oxidative fermentation but also to other fermentation industries, since a huge amount of expense is required for cooling. A temperature increase by 2–3 ºC causes a serious failure in both fermentation rate and fermentation efficiency. In submerged cultures, a large amount of heat is generated during fermentation and cooling costs become


even more expensive. If favorable strains of acetic acid bacteria that can work optimally at 37–40 ºC were available, such strains would be able to accept loose temperature control and the cooling expenses would be reduced considerably. The thermotolerant strain of G. frateurii CHM 54 was able to produce L-sorbose at higher temperatures. When D -sorbitol oxidation was done at higher temperatures with the thermotolerant strain, both the fermentation rate as well as fermentation efficiency were superior to that achieved using non-thermotolerant mesophilic strains [66].

1.3.3.8

L-Sorbosone

L-Sorbosone


→ L-ascorbate

In current industrial L-ascorbate production processes, 2-keto-L-gulonate is a key intermediate that is chemically converted to L-ascorbate. All processes known to date require a large amount of energy and organic solvent, and thus a cheaper and environmentally conscious substitute process, such as enzymatic conversion, is being looked for. Due to the practical importance of the enzymes leading to Lascorbate production, there are many papers dealing with the enzymes concerned (see Fig. 1.7). The following outstanding results should be itemized: isolation and characterization of a new PQQ-dependent dehydrogenase, L-sorbose/L-sorbosone dehydrogenase [70], cloning the genes coding for L-sorbose and L-sorbosone dehydrogenases from Gluconobacter oxydans and microbial production of 2-ketoL-gulonate, a precursor of L-ascorbate, in a recombinant G. oxydans [71], cloning and nucleotide sequencing of the membrane-bound L-sorbosone dehydrogenase gene of Acetobacter liquefaciens IFO 12258 and its expression in Gluconobacter oxydans [72], isolation and characterization of a new vitamin C-producing enzyme (L-gulono-γ-lactone dehydrogenase of bacterial origin [73], microbial producton of L-ascorbate from D -sorbitol, L-sorbose, L-gulose, and L-sorbosone by Ketogulonicienium vulgare DSM 4025 [74]. Recently, a novel PQQ-dependent enzyme, L-sorbosone dehydrogenase 1 (SNDH1) catalyzing direct conversion of L-sorbosone to L-ascorbate was purified from Ketogluconicigenium vulgare DSM 4025 [75]. SNDH1 is a homooligomer of 75 kDa subunits containing PQQ and heme c as the prosthetic group. Two isozymes of SNDH1, SNDH2 consisting of 75 kDa and 55 kDa subunits, and SNDH3 consisting of a 55 kDa subunit, were also purified from the same strain. It was found that the 55 kDa subunit was derived from the 75 kDa subunit after cleavage of the C-terminal domain in the bacterial cells. The three enzymes catalyzed L-ascorbic acid formation as well as 2-keto-L-gulonate from L-sorbose, suggesting that tautomerization of L-sorbosone causes the dual conversion by SNDHs. Industrial L-ascorbic acid production has revealed direct conversion of L-sorbosone to L-ascorbic acid by a membrane-bound quinoprotein L-sorbosone dehydrogenase.

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1.3.4 Quinate Oxidation. Membrane-Bound Quinate Dehydrogenase (QDH)

Quinate → 3-dehydroquinate Unlike enzymes relating to the shikimate pathway found in the cytoplasm of microorganisms, the first report of a quinate-oxidizing enzyme in acetic acid bacteria was done by Whiting and Coggins [76]. They described quinate oxidation to 3-dehydroquinate (DQA) by an NAD(P)-independent quinate dehydrogenase (QDH) (EC 1.1.99.25) and shikimate (SKA) oxidation to 3-dehydroshikimate (DSA) by an enzyme associated with the particulate enzyme in the cytoplasmic membrane. van Kleef and Duine described the occurrence of QDH in the periplasm of Acinetobacter calcoaceticus LMD 79.41 and suggested that QDH is a quinprotein in which PQQ is involved [77] (Fig. 1.8). QDH purification was done with Gluconobacter oxydans IFO 3244 and Acinetobacter calcoaceticus AC3 after solubilization with detergent [78, 79]. Due to the hydrophobicity of QDH as a typical membrane-bound enzyme, QDH is one of the most difficult enzymes to purify to high homogeneity. The molecular mass of QDH was estimated to be 88 kDa. It oxidizes quinate and SKA with optimal activity at pH 6–7. Since A. calcoaceticus AC3 cannot produce PQQ, QDH was

Fig. 1.8 Metabolic map of shikimate pathway.


purified as an apoenzyme consisting of dimeric structure, which was converted to the monomeric holoenzyme on addition of PQQ. QDH is very beneficial enzyme for the production of SKA via DQA and DSA from quinate supplied outside the SKA pathway without any metabolic control repressing SKA production. In 2003, we proposed SKA production by a single cellular system of acetic acid bacteria [80–83]. QDH and 3-dehydroquinate dehydratase (DQD) are located predominantly on the outer surface of the cytoplasmic membranes of some species of Gluconobacter strains and quinate is oxidized to DSA via DQA in a sequential manner. In the cytoplasm, SKDH catalyzes a reversible reaction of SKA oxidation to DSA and DSA reduction to SKA. However, many trial and error experiments to combine together the two separately located enzymatic systems taking place outside and inside the cells only resulted in insufficient production of SKA. As a more positive response to the global need for oseltamivir, we have developed a better strategy for high SKA production, allowing the two separately located enzymatic systems to work sequentially. Dried cells or the membrane fraction involving QDH and DQD may be used for DSA production in the first reaction (system 1 in Fig. 1.9a). The second reaction is a coupling reaction composed of two cytosolic enzymes, SKDH and GDH, as an NADPH-regenerating enzyme (system 2 in Fig. 1.9a). The coupling reaction by the two cytosolic enzymes in the presence of excess D -glucose works well as expected until the DSA added initially is converted completely to SKA. The overall reaction carried out for SKA production by the two different enzymatic systems is depicted in Fig. 1.9b. SKA is a key intermediate for aromatic amino acids as well as for large numbers of antibiotics, alkaloids, and herbicides. Recently, another important role for SKA has emerged as a precursor for the synthesis of oseltamivir (Tamiflu), an antiviral drug designed to protect people from pandemic flu infection. In spite of warnings from the World Health Organization about the approach of a global flu pandemic, including avian influenza, there are insufficient stocks of oseltamivir around the world. One reason for this is the technical difficulties in preparing SKA, because two different metabolic pathways, glycolysis and pentose phosphate pathway, need to be combined before forming SKA. Furthermore the metabolic location of SKA is a long way from that for glucose and it is very difficult to lead the metabolic flow to SKA production through classic fermentation technology as well as through modern molecular biotechnology. Although resources are limited, SKA is also produced by extraction from plants such as Illicium anisatum or I. verum. The total synthesis of SKA through organic chemistry has not been practically available. Nevertheless, we need to address the challenge of developing a novel method for more effective and convenient SKA production. SKA is remote from D -glucose in the metabolic pathway, the more shortcut access to the SKA pathway from quinate looks advantageous to produce the important metabolic intermediates generated in the SKA pathway. DSA is formed with a yield of about 90% from quinate via DQA by two suc-cessive enzyme reactions, QDH and 3-dehydroquinate dehydratase (DQD) (EC 4.2.1.10) located in the

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Fig. 1.9 (a) Overall reaction for SKA production and (b) enzymatic conversion of DSA to SKA with SKDH coupled with GDH.

cytoplasmic membranes of acetic acid bacteria. DSA is then reduced to SKA with NADP-dependent SKA dehydrogenase (SKDH) (EC 1.1.1.25) from the same organism. When SKDH is coupled with NADP-dependent glucose dehydrogenase (GDH) (EC 1.1.1.47) in the presence of excess D -glucose as an NADPH regenerating system, SKDH works to produce SKA until DSA added initially in the reaction mixture is completely converted to SKA as shown in Fig. 1.9b [83, 84]. For the system 1, the dried cells or the membrane fraction were incubated with quinate at pH 5.0 overnight with shaking. The DSA formed was measured with the deproteinized supernatants. Since DQD have a pH optimum at 8.0 and low activity of DQD is found at pH 5.0, the reaction can be regulated to make DQA as the major product. The apparent conversion rate from quinate to DQA is estimated at over 90%. On the other hand, when the above reaction was carried out

1.4 FAD-Dependent Dehydrogenase

at pH 8.0, DSA was accumulated as the major product, because DQA formed can be converted immediately to DSA under the optimum condition of DQD. The apparent yield of DSA from quinate is usually over 90% [85]. Thus, a convincing strategy for preparing commercially unavailable metabolic intermediates, DQA and DSA, by means of bioconversion has been established. QDA is one of the compounds involved in the oxidative fermentation, the initially added quinate is converted to the corresponding oxidation product with high yield, as seen with other PQQ- and FAD-dependent dehydrogenases. A simple chromatographic method for separating the four compounds – quinate, DQA, DSA, and SKA – in the SKA pathway has recently become available [85].

1.4 FAD-Dependent Dehydrogenase 1.4.1 D -Fructose Dehydrogenase, Membrane-Bound D -Fructose D -Fructose

→ 5-keto- D -fructose

oxidation to 5-keto- D -fructose (5KF) is catalyzed by membrane-bound fructose dehydrogenase (FDH, EC 1.1.99.11), which contains a covalently bound FAD as the primary coenzyme. FDH was purified from the membrane fraction of G. industrius IFO 3260. FDH was solubilized from the membrane with 1% Triton X-100 and further purified to a homogeneous state [86]. The purified FDH had a sedimentation coefficient of 5.8S and the total molecular mass was estimated to be 140 kDa by gel fi ltration. The enzyme was dissociated into three different subunits, 67 kDa, 50 kDa, and 19 kDa, during gel electrophoresis. The largest subunit was positive to enzyme activity staining in a mixture containing PMS, nitroblue tetrazolium, and D -fructose. When an unstained gel was irradiated with a fluorescent light, intense fluorescence was observed only with the largest subunit, indicating the presence of a covalently bound FAD as the coenzyme. The second subunit (50 kDa) was characterized as the subunit carrying cytochrome c. Regarding the smallest subunit, as with ADH III, GADH, KGDH, and other membrane-bound dehydrogenases, the actual function and properties of the subunit were unknown. However, if the smallest subunit is deleted, the enzyme activity of the individual enzymes is also halted and no accumulation of the corresponding oxidation product outside the cells is observed. Thus, the smallest subunit is essential to enzyme activity as well as to the oxidative fermentation. It should be noted that only D -fructose was oxidized by FDH during the study of substrate specificity with various substrates. When D -fructose was oxidized in the presence of the same concentration of substrate analogs such as D -glucose, D -mannose, D -fructose-6-phosphate, D -fructose-1,6-diphosphate, 5-keto- D fructose, D -glucose-6-phosphate, D -glucose-1-phosphate, D -gluconate, 2-keto- D -

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gluconate, and 5-keto- D -gluconate, the reaction rate of D -fructose oxidation was not affected at all. These properties of FDH suggest that FDH could be beneficial as the enzyme for D -fructose microdetermination. As mentioned below, an apparent Michaelis constant determined at pH 4.5 was found to be 10 mmol L −1. The optimum pH for D -fructose oxidation was found at pH 4.0–4.5. The reaction product of D -fructose oxidation was identified to be 5-keto- D -fructose, which has no negative effect on D -fructose oxidation. As has already mentioned above, the most characteristic point of oxidative fermentation is that D -fructose oxidation by FDH continues until the D -fructose added initially in the reaction mixture is oxidized completely to the reaction product; there is no reaction equilibrium, unlike NAD(P)-dependent dehydrogenases. With respect to the enzymatic method for D -fructose determination, a coupling enzymatic method involving hexokinase (EC 2.7.11), phospho- D -glucose isomerase ( D -glucose-6-phosphate ketol isomerase, EC 5.3.1.9), and D -glucose-6phosphate dehydrogenase (EC 1.1.1.49) is the sole reliable method so far reported [87]. However, this assay method depends largely on the purity of the enzymes used and therefore is rather expensive. Difficulty in preparing the required enzymes without any contaminants could make this assay system complicated and potentially troublesome. A trace of D -glucose present in the samples for D -fructose assay invites inevitable confusion, because hexokinases react widely with many aldoses. The preparation of highly purified D -fructokinase that reacts with only D -fructose is almost impossible. Hence, the use of FDH is highly recommended instead of the method above, because FDH reacts only with D -fructose so can be used for the rate assay as well as for the end point measurement. In this chapter, it is emphasized that PQQ- and FAD-dependent dehydrogenases obtained from the membrane fraction of acetic acid bacteria and other aerobic bacteria are useful and convenient enzymes for biosensors monitoring ethanol, acetaldehyde, D -glucose, D -gluconate, 2-keto- D -gluconate, and so on. In the same way, FDH can be the enzyme for D -fructose measurement for clinical purposes [88]. FDH could be beneficial for agriculture, for example in fruit production to check the best timing for ripening, and in fermentation industries for process control in brewing. FDH is now available on the market.

1.4.2 D -Gluconate Dehydrogenase, Membrane-Bound D -Gluconate D -Gluconate

→ 2-keto- D -gluconate

dehydrogenase (GADH) (EC 1.1.99.3) occurs on the outer surface of cytoplasmic membranes of aerobic bacteria, such as Pseudomonas, Klebsiella, Serratia, and acetic acid bacteria. The enzyme activity is linked to the electron transport chain in the cytoplasmic membrane constituting a D -gluconate oxidase system [89–91]. GADH of P. aeruginosa shows a single protein band on native PAGE, but the enzyme preparations from K. klebsiella and S. marcescens are separated into two


protein bands [92]. The low-mobility band is yellow and fluorescent under ultraviolet light, and shows enzyme activity. The coenzyme functioning has been shown to be 8α-[N3-histidyl)riboflavin from the largest subunit [93]. Another protein band with higher mobility is red-colored. In SDS-PAGE, enzyme preparations from four different bacterial strains dissociate into three different polypeptide bands, subunit I having 66–68 kDa, subunit II (48–52 kDa), and subunit III (22–25 kDa). The sum of the three subunits gives 130–140 kDa as the total molecular mass. GADH is a monomeric protein dispersed in the presence of 0.1% detergent such as Triton X-100, whereas the deletion of detergent allows GADH to be the dimeric enzyme. The removal of detergent from the purified enzyme causes a decrease of enzyme activity. Activation of GADH is observed with phospholipids, especially cardiolipin, in the presence of Triton X-100. Thus, GADH has a hydrophobic and phospholipid-interacting domain, and hence is a typical membrane-bound dehydrogenase. All GADHs are highly specific for D -gluconate oxidation with K m values of 0.3–0.8 mmol L −1 and show optimum pH at 4.0–5.0, when assayed with potassium ferricyanide. The following compounds are not oxidized by GADH: aldohexoses, aldopentoses, 2-keto- D -gluconate, 5-keto- D -gluconate, D -galactonate, D -mannonate, 6-phospho- D -gluconate, L-idonate, D -arabonate, and D -xylonate. Since the enzyme activity of GADH is not affected at all by the presence of high concentrations of the above compounds, GADH is a favorable enzyme for D -gluconate measurement [94]. D -Gluconate is well known as one of the natural ingredients of brewing products and is widespread in other materials as food additives. In spite of the wide distribution and usefulness of D -gluconate, an accurate method for its quantitative measurement had not been developed until GADH was proposed for the purpose. A well-known coupling enzymatic D -gluconate determination developed and used so far contains D -gluconate kinase and 6phospho- D -gluconate dehydrogenase. However, this assay method depends largely on the purity of the enzymes used and is rather expensive. Difficulty in preparing a pure D -gluconate kinase and 6-phospho-D -gluconate dehydrogenase without any contaminants makes this assay system complicated and troublesome. A simple chemical colorimetric determination using GADH from aerobic bacteria is highly reliable method because GADH is prepared with ease and the enzyme activity is stable without appreciable loss if GADH is stored in an appropriate buffer solutions in the presence of detergent. Rate assay as well as the end point measurement for D -gluconate can be done successfully with GADH. 1.4.3 D -Hexosamine Dehydrogenase, Membrane-Bound

→ D -glucosaminate → D -galactosaminate D -Mannosamine → D -mannosaminate D -Glucosamine

D -Galactosamine

In most strains of acetic acid bacteria, membrane-bound D -hexosamine dehydrogenase (tentatively named) oxidizes D -hexosamines to the corresponding D -

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hexosaminates stoichiometrically. Conversion of D -hexosamines into D -hexosaminates is observed with growing cells of acetic acid bacteria and D -hexosaminate is accumulated in the culture medium after D -hexosamine is exhausted. Since the responsible enzyme is accommodated on the outer surface of the cytoplasmic membrane and the enzyme activity is linked to the respiratory chain of the organisms, resting cells, dried cells, and immobilized cells of acetic acid bacteria are effective catalyst for D -hexosaminate production. D -Mannosaminate and D -galactosaminate, commercially unavailable compounds, can be prepared with ease with acetic acid bacteria. The respective three different D -hexosaminates are shown to be separated from each other by chromatography [95]. The first report of D -glucosaminate formation by acetic acid bacteria and other aerobic bacteria like pseudomonads was carried out in experiments with resting cells and D -glucosaminate was indirectly suggested to be formed as the oxidation product of D -glucosamine [96]. Recently, bioconversion of D -glucosamine to D glucosaminate by a strain of Acinetobacter sp. isolated by enrichment techniques on D -glucosamine was briefly reported [97]. Although acetic acid bacteria are different from Acinetobacter in many respects, the enzyme functioning in D glucosamine oxidation in Acinetobacter sp. is supposed to be the same as in acetic acid bacteria. There is no information about the substrate specificity of quinoprotein D -glucose dehydrogenase (m-GDH) or whether the enzyme is capable of D -glucosamine oxidation. The enzyme activity of D -glucosamine dehydrogenase can be measured using either a combination of PMS and DCIP assayed in 50 mmol L −1 potassium phosphate, pH 7.0, or potassium ferricyanide assayed in 50 mmol L −1 glycine-NaOH buffer, pH 9.0, under essentially the same conditions as described by Ameyama [3]. Identification and measurement of D -hexosaminate can be done enzymatically with D -glucosaminate dehydratase (EC 4.2.1.26) purified from the cell-free extract of P. fluorescens IFO 14808 according to Iwamoto and Imanaga [98]. When the growth profi le of G. frateurii IFO 3264 with high D -hexosamine oxidase activity was examined in 0.5% D -glucosamine medium and monitored both by measuring turbidity and by direct viable cell counting, the organism survived even though cultivation was prolonged to the late stationary phase, where the D glucosamine was exhausted and converted to D -glucosaminate [95]. Unlike the case of Acinetobacter sp. [97], it is characteristic to see that the majority of the cells of G. frateurii IFO 3264 survived over the prolonged cultivation after complete oxidation of D -glucosamine to D -glucosaminate, as demonstrated by viable cell counting throughout the cultivation. D -Glucosamine initially added to the culture medium was converted to D -glucosaminate almost stoichiometrically, indicating that the oxidation products of D -glucosamine stayed stable without any significant breakdown by the organism. This is a noticeable difference from the case of pseudomonads or other Gram-negative bacteria, in which an appreciable amount of D -glucosaminate was further assimilated, as also suggested by Takahashi and Kayamori [96]. Encouraged by the clear data on D -hexosamine oxidation as shown above, workers have examined the microbial conversion of D -mannosamine and D -


galactosamine to yield corresponding D -hexosaminates [95]. Freshly harvested cells (10 mg mL −1) of G. frateurii IFO 3264 grown to the stationary phase were mixed with either D -mannosamine-HCl or D -galactosamine-HCl in 5 mmol L −1 potassium phosphate, pH 7.0, and the total volume was adjusted to 10 mL. The mixtures were kept at 30 ºC with shaking and the remaining D -hexosamine in the reaction mixture was checked periodically with a purified D -glucosamine dehydrogenase. D -Mannosamine oxidation took 2 h and D -galactosamine oxidation took 8 h under the above conditions. As additional new information, D -glucosaminate dehydratase was just as effective as the three different D -hexosaminates. For example, 136% and 118% of the relative reaction rate to D -glucosaminate were observed for D -mannosaminate and D -galactosamainate, respectively, when individual oxidation products were assayed with D -glucosaminate dehydratase under steady state conditions. Thus, production of D -mannosaminate and D galactosaminate was indicated successfully for the first time, accompanied by the enzymatic identification of the individual oxidation products. We have to mention the reason why D -hexosamine oxidizing enzyme is classified as an FAD-dependent dehydrogenase. The main reason, after solubilization and partial purification of the respective enzyme, is that potassium ferricyanide was still valid as the electron acceptor. Second, the solubilized enzyme was highly resistant to EDTA treatment indicating either that it contains a covalently bound FAD or that the PQQ binding to the enzyme is unusually strong. The fi nal answers will be found soon. 1.4.4 2-Keto- D -gluconate Dehydrogenase, Membrane-Bound

2-Keto- D -gluconate → 2,5-diketo-D -gluconate The enzyme catalyzing 2-ketogluconate oxidation, yielding 2,5-diketo-D gluconate, has been characterized as a flavohemoprotein with three different subunits [99]. From the membrane fraction of Gluconobacter melanogenus IFO 3293, 2-keto- D -gluconate dehydrogenase (KGDH) is solubilized and purified with high yield to a homogeneous state examined by the criteria of electrophoresis and analytical ultracentrifugation. The purified KGDH is homogeneous in analytical ultracentrifugation, with a sedimentation coefficient of 5.9 S, and also in nativePAGE, with a single protein band. After solubilization of the enzyme performed with 2% Na-cholate and 0.2 mol L −1 KCl, two-step column chromatography gave a purified enzyme. The molecular mass was measured to be 133 kDa and SDSPAGE shows the presence of three different subunits of 61 kDa (flavoprotein), 47 kDa (cytochrome c), and 25 kDa. The flavoprotein contained a covalently bound FAD [93]. Purified KGDH has a characteristic deep rose-red color due to the cytochome component. The typical reduced cytochrome c type absorption spectrum has maxima at 554, 523, and 417 nm. The cytochrome component of the enzyme is reduced on addition of either 2-keto- D -gluconate or sodium dithionite. A

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successful purification of KGDH came from the finding of fundamental differences in hydrophobicity between D -gluconate dehydrogenase (GADH) (see Section 1.4.2) and KGDH. KGDH was not solubilized in the absence of a chaotropic agent, KCl. With Brij 35 and Tween 80, even in the presence of KCl, KGDH was scarcely solubilized, but GADH was solubilized with a final recovery of 100%. Thus, the membrane fraction of G. melanogenus IFO 3293 was treated first with 2% Brij 35 and 0.2 mol L −1 KCl to remove GADH, and after centrifugation the precipitate containing KGDH was collected. The resulting membrane precipitates were suspended in 0.01 mol L −1 Tris-HCl, pH 8.0, and treated with Triton X-100, Na-cholate, or Na-deoxycholate for 3 h at 5 ºC. KGDH was solubilized in the presence of 0.3 mol L −1 KCl together with 2% Triton X-100. Alternatively, solubilization of KGDH was carried out overnight with 2% Na-cholate and 0.2 mol L −1 KCl, which was favorable to the following purification procedure involving ammonium sulfate fractionation. The enzyme activity of KGDH is most active at pH 4.0, and 2-keto- D -gluconate is the only substrate oxidized by the enzyme. Similar compounds such as 5-ketoD -gluconate, 2-keto- D -galactonate, and 2-keto- D -gulonate are inert. KGDH catalyzes 2-keto- D -gluconate oxidation to 2,5-diketo-D -gluconate, which is an important step of bioconversion in a novel pathway to L-ascorbate developed by Sonoyama et al. [100]. D -Glucose was first converted to Ca-2,5-diketo-D -gluconate by a mutant strain of Erwinia sp. with 94.5% yield after 26 h cultivation. Then, a mutant strain of Coprynebacterium sp. reduced 2,5-diketo- D -gluconate stereospecifically with 2,5-diketo- D -gluconate reductase to Ca-2-keto-L-gulonate with a yield of 84.5%. The occurrence of KGDH is known in strains of Erwinia, Pantoea, and Pseudomonas, as well as Gluconobacter. Ameyama and Kondo noted the importance of 2,5-diketo- D -gluconate as the precursor of D -lyxuronic acid [101]. 1.4.5 Sorbitol Dehydrogenase, Membrane-Bound

Sorbitol → L-sorbose Unlike the ubiquitous quinoprotein polyol dehydrogenases in acetic acid bacteria, one of which catalyzes oxidation of D -sorbitol to L-sorbose, a distinct D -sorbitol dehydrogenase (SLDH) was found in the membrane fraction of G. suboxydans var. a IFO 3254. SLDH was solubilized from the membrane fraction and purified as a covalently bound FAD-containing dehydrogenase [67]. The best solubilization of SLDH carried out in 10 mmol L −1 acetate, pH 5.0, containing 1% Triton X-100, 0.1 mol L −1 KCl, and 0.1 mol L −1 D -sorbitol gave the highest recovery of more than 100%. Purification of SLDH was readily done by DEAE-cellulose and CMcellulose chromatography. The purified SLDH showed a rose-red color due to the presence of heme c and the absorption spectrum of SLDH after reduction with sodium dithionite gave absorption maxima at 551, 522, and 417 nm in the visible region. The heme c component was not reduced rapidly by the addition of D sorbitol, primary substrate, until coenzyme Q1 was added to the enzyme solution,

1.5 Miscellaneous

implying that ubiquinone mediates D -sorbitol oxidation, similar to the other membrane-bound dehydrogenases exemplified in this article. SLDH dissociated into three subunits of 63 kDa (subunit I), 51 kDa (subunit II), and 17 kDa (subunit III). Subunit I was characterized as a flavoprotein and FAD was bound covalently to the enzyme. Subunit II contained heme c. SLDH catalyzes D -sorbitol oxidation to L-sorbose exclusively, which is distinct from PQQ-dependent polyol dehydrogenases catalyzing D -sorbitol oxidation to Lsorbose. The substrate specificity is restricted to D -sorbitol, and D -mannitol was oxidized only by 5% of that of D -sorbitol. Other polyols, such as D -arabitol, D -iditol, meso-erythritol, dulcitol, ribitol, and xylitol, are not oxidized by the enzyme.

1.5 Miscellaneous 1.5.1 Aldehyde Dehydrogenase, Membrane-Bound

Aldehyde → Carboxylic acid Membrane-bound aldehyde dehydrogenase (ALDH) in acetic acid bacteria acts on a wide range of aliphatic aldehydes except for formaldehyde. Aldehydes with a carbon chain length of 2–4 are oxidized most rapidly with ALDH from both genera of Acetobacter and Gluconobacter. The enzyme is localized on the outer surface of the cytoplasmic membrane of the organisms and has a close topological and functional relation to ADH III. Aldehyde oxidation is linked to the respiratory chain as described for the alcohol oxidase system above. Thus, ALDH acts as vinegar producer sequentially after ADH III in acetic acid bacteria. During alcohol oxidation, no aldehyde liberation is observed under normal culture conditions, indicating that ADH III and ALDH form a multienzyme complex in the bacterial membrane and function sequentially to produce acetate from ethanol. As for ALDH from acetic acid bacteria, the purification and characterization of ALDH have been done with several strains [102–106]. As summarized by Matsushita et al. [1], purified ALDHs from G. suboxydans and A. aceti contain heme component, while ALDHs from A. polyoxogenes and A. rancens do not contain heme c. The subunit structures and compositions from the different sources are different. A composition of two subunits was reported, with ALDH from G. suboxydans composed of subunit I (86 kDa) and subunit II (55 kDa), A. polyoxogenes, subunit I (75 kDa) and subunit II (19 kDa), and A. rancens, subunit I (78 kDa) and subunit II (66 kDa). ALDH from A. aceti is composed of threee subunits: subunit I (78 kDa), subunit II (45 kDa), and subunit III (14 kDa). The subunit I contains the catalytic site involving a molybdoputerin cofactor as the primary coenzyme. Purified ALDHs containing heme c from G. suboxydans [102] and A. aceti [103] are rose-red in color with absorption maxima at 551, 523, and 418 nm (reduced

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enzyme) and a sole peak at 410 nm (oxidized enzyme). Since the absorption spectra of ALDH is very similar to that of ADH III, ALDH was believed to be a PQQ-dependent enzyme. However, a mutant lacking the gene encoding PQQ biosynthesis still contained active ALDH, while the enzyme activity of ADH III was completely lost, indicating that the coenzyme of ALDH is not PQQ [107]. Following cloning of an ALDH-encoding gene from A. europaeus the deduced amino acid sequence indicated the presence of a molybdenum–molybdopterin cytosine dinucleotide coenzyme [108]. Molybdenum–molybdopterin cytosine dinucleotide was also indicated as the cofactor of isoquinoline 1-oxidoreductase of Pseudomonas diminuta 7 [109]. ALDH may have a similar coenzyme structure, although the final characterization of the coenzyme in ALDH has not been completed. ALDH from A. aceti is highly stable in acidic pH as well as highly resistant to heating and more than 50% of aldehyde oxidase in the membrane fraction survived when heated at 60 ºC for 30 min, while enzyme activity of ADH III was lost rapidly within a few minutes [110]. A biocatalyst composed of the ALDH-containing membranes of acetic acid bacteria is useful in eliminating off-flavors caused by various middle chain length aliphatic aldehydes that occur in foodstuffs such as wheat flour and soybean meal. When unripe cereal grains or beans are used in flour making or soybean meal production, strong off-flavors cause serious problems. Most aldehydes have a low threshold of off-flavor but once such aldehydes are oxidized to the corresponding carboxylic acids, for which the threshold is quite high, the off-flavors are decreased. Like alcohol yeasts and lactic acid bacteria, acetic acid bacteria are known to be edible microbes and produce no appreciable pathogens. Therefore, there is no problem in using cells of acetic acid bacteria carrying ALDH to improve the quality of foodstuffs. The membrane fraction containing ALDH exclusively produced by deleting ADH III by heating is also a useful enzyme for aldehyde microdetermination [110]. As for NAD(P)-independent formaldehyde dehydrogenase, the enzyme from Methylococcus capsulatus Bath purified from the membrane was shown to have PQQ as a cofactor [111]. This is the first report of the purification of NAD(P)independent formaldehyde dehydrogenase from the membrane fraction of a methylotroph and PQQ-containing formaldehyde dehydrogenase coupled to the electron transport chain via a β-type cytochrome or quinone. The properties of this enzymes showed a number of similarities to the soluble NAD(P)independent ALDH from Hyphomicrobium zavarzinii ZV 580 [112]. Another formaldehyde-oxidizing enzyme is the quinoprotein methanol dehydrogenase from methylotrophic and methanotrophic bacteria [2, 113]. So far, formaldehyde dehydrogenase, including both NAD(P)-dependent and NAD(P)independent enzymes, has been studied mainly in bacteria and yeasts that are able to grow on C1 compounds such as methane and methanol as sole carbon source. It is interesting to see that formaldehyde oxidizing ADH III has recently been found in Acetobacter sp. [114].

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dehydrogenase is independent of the oxidative fermentation. Biosci Biotechnol Biochem 2001, 65, 115–125. Adachi, O., Fujii, Y., Ghaly, F.M., Toyama, H., Shinagawa, E., Matsushita, K. Membrane-bound quinoprotein D arabitol dehydrogenase of Gluconobacter suboxydans IFO 3257, A versatile enzyme for the oxidative fermentation of various ketoses. Biosci Biotechnol Biochem 2001, 65, 2755–2762. Hann, M.R., Tilden, B.E., Hudson, C.S. The oxidation of sugar alcohols by Acetobacter suboxydans. J Am Chem Soc 1938, 60, 1201–1203. Ameyama, M., Shinagawa, E., Matsushita, K., Adachi, O. Solubilization, purification and properties of membrane-bound glycerol dehydrogenase from Gluconobacter industrius. Agric Biol Chem 1985, 49, 1001–1010. Sugisawa, T., Hoshino, T. Purification and properties of membrane-bound D -sorbitol dehydrogenase from Gluconobacter suboxydans IFO 3255. Biosci Biotechnol Biochem 2002, 66, 57–64. Ameyama, M., Adachi, O. 5-Keto- D gluconate reductase from Gluconobacter suboxydans. Methods Enzymol 1982, 89, 198–202. Ameyama, M., Adachi, O . 2-Keto- D gluconate reductase from acetic acid bacteria. Methods Enzymol 1982, 89, 203–210. Bertrand, G. Novel preparation of L-erythrulose by meso-erythritol dehydrogenation. C R Seances Acad Sci 1900, 130, 1472–1475. Whistler, R.L., Underkofler, L.A. The production of L-erythrulose by the action of Acetobacter suboxydans upon erythritol. J Am Chem Soc 1938, 60, 2507–2508. Moonmangmee, D., Shinagawa, E., Toyama, H., Theeragool, G., Lotong, N., Matsushita, K., Adachi, O. L-Erythrulose production by oxidative fermentation is catalyzed by PQQcontaining membrane-bound dehydrogenase. Biosci Biotechnol Biochem 2002, 66, 307–318.

60 Asai, T. Oxidation of monosaccharides. In: Asai, T., ed. Acetic Acid Bacteria. Tokyo: University of Tokyo Press, 1968, pp. 178–203. 61 Matsushita, K., Fujii, K., Ano, Y., Toyama, H., Shinjoh, M., Tomiyama, N., Miyazaki, T., Hoshino, T., Adachi, O. 5Keto- D -gluconate production is catalyzed by a quinoprotein glycerol dehydrogenase, major polyol dehydrogenase, in Gluconobacter sp. Appl Environ Microbiol 2003, 69, 1959–1966. 62 Gray, G.E. Preparation of 2-keto gulonic acid and its salts. US Patent 2,421,611, 1945. 63 Gray, G.E. Preparation of 2-keto gulonic acid and its salts. US Patent 2,421,612, 1945. 64 Fulmer, E.I., Dunning, J.W., Underkofler, L.A. The effect of the concentration of mannitol upon the production of levulose by the action of Acetobacter suoxydans. Iowa State Coll J Sci 1939, 13, 279–281. 65 Arcus, A.C., Edson, N.L. Polyol dehydrogenases. 2. The polyol dehydrogenases of Acetobacter suboxydans and Candida utilis. Biochem J 1956, 64, 385–394. 66 Moonmangmee, D., Adachi, O., Ano, Y., Shinagawa, E., Toyama, H., Theeragool, G., Lotong, N., Matsushita, K. Isolation and characterization of thermotolerant Gluconobacter strains catalyzing oxidative fermentation at higher temperatures. Biosci Biotechnol Biochem 2000, 64, 2306–2315. 67 Shinagawa, E., Matsushita, K., Adachi, O., Ameyam, M. Purification and characterization of D -sorbitol dehydrogenase from membrane of Gluconobacter suboxydans var. α. Agric Biol Chem 1982, 46, 135–141. 68 Adachi, O., Toyama, H., Theeragool, G., Lotong, N., Matsushita, K. Crystallization and properties of NAD-dependent D sorbitol dehydrogenase from Gluconobacter suboxydans IFO 3257, Biosci Biotechnol Biochem 1999, 63, 1589–1595. 69 Adachi, O., Ano, Y., Moonmangmee, D., Shinagawa, E., Toyama, H., Theeragool,

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by Acetomonas oxydans. Biochem J 1967, 102, 283–293. van Kleef, M.A.G., Duine, J.A. Bacterial NAD(P)-independent quinate dehydrogenase is a quinoprotein. Arch Miocrobiol 1988, 150, 32–36. Adachi, O., Yoshihara, N., Tanasupawat, S., Toyama, H., Matsushita, K. Purification and characterization of membrane-bound quinoprotein quinate dehydrogenase. Biosci Biotechnol Biochem 2003, 67, 2115–2123. Vangnai, A.S., Toyama, H., Deeknamkul, W., Yoshihara, N., Adachi, O., Matsushita, K. Quinate oxidation in Gluconobacter oxydans IFO3244: purification and characterization of quinoprotein quinate dehydrogenase. FEMS Microbiol Lett 2004, 241, 157–162. Adachi, O., Moonmangmee, D., Toyama, H., Yamada, M., Shinagawa, E., Matsushita, K. New quinoproteins in oxidative fermentation. Biochim Biophys Acta 2003, 1647, 10–17. Adachi, O., Moonmangmee, D., Toyama, H., Yamada, M., Shinagawa, E., Matsushita, K. New developments in oxidative fermentation. Appl Microbiol Biotechnol 2003, 60, 643–653. Adachi, O., Tanasupawat, S., Yoshihara, N., Toyama, H., Matsushita, K. 3Dehydroquinate production by oxidative fermentation and further conversion of 3-dehydroquinate to the intermediates in the shikimate pathway. Biosci Biotechnol Biochem 2003, 67, 2124–2131. Adachi, O., Ano, Y., Toyama, H., Matsushita, K. High shikimate production from quinate with two enzymatic systems of acetic acid bacteria. Biosci Biotechnol Biochem 2006, 70, 2579–2582. Adachi, O., Ano, Y., Toyama, H., Matsushita, K. Purification and properties of shikimate dehydrogenase from Gluconobacter oxydans IFO 3244 and its application to enzymatic shikimate production. Biosci Biotechnol Biochem 2006, 70, 2786–2789. Adachi, O., Ano, Y., Toyama, H., Matsushita, K., Enzymatic preparation of metabolic intermediates, 3-dehydroquinate and

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3-dehydroshikimate, in the shikimate 94 Ameyama, M., Tayama, K., Shinagawa, pathway. Biosci Biotechnol Biochem E., Matsushita, K., Adachi, O. New 2006, 70, 3081–3083. enzymatic determination of gluconate Ameyama, M., Shinagawa, E., with particulate D -gluconate Matsushita, K., Adachi, O. D -Fructose dehydrogenase. Agric Biol Chem 1978, dehydrogenase of Gluconobacter 42, 2347–2354. industrius: Purification, 95 Moonmangmee, D., Adachi, O., Toyama, characterization and application to H., Matsushita, K. D -Hexosaminate enzymatic microdetermination of D production by oxidative fermentation. fructose. J Bacteriol 1981, 145, 814–823. Appl Biotechnol Microbiol 2004, 66, Bent, E., Bergmeyer, H.U. D -Fructose. 253–258. Methods Enzymatic Anal 1974, 96 Takahashi, K., Kayamori, H. Studies on 3, 1304–1307. the formation of glucosaminic acid by Nakashima, K., Takei, H., Adachi, O., Acetobacter melanogenum Beijerinck Shinagawa, E., Ameyama, M. and Pseudomonas fluorescens. Bull Determination of seminal fructose Agric Chem Soc Jpn 1960, 24, using D -fructose dehydrogenase. Clin 231–234. Chim Acta 1985, 151, 307–310. 97 Takiguchi, Y., Shiina, A., Yamaguchi, Matsushita, K., Shinagawa, E., Adachi, T.T. Bioconversion of D -glucosamine O., Ameyama, M. Membrane-bound to D -glucosaminic acid. Nippon D -gluconate dehydrogenase from Nogeikagaku Kaishi 2003, 77, 576–578 Pseudomonas aeruginosa. Purification (in Japanese). and structure of cytochrome-binding 98 Iwamoto, R., Imanaga, Y., Purification form. J Biochem 1979, 85, 1173–1181. and characterization of D -glucosaminate Matsushita, K., Shinagawa, E., Adachi, dehydratase from Pseudomonas O., Ameyama, M. Membrane-bound fluorescens. Agric Biol Chem 1989, 53, D -gluconate dehydrogenase from 2563–2569. Pseudomonas aeruginosa. Its kinetic 99 Shinagawa, E., Matsushita, K., Adachi, properties and a reconstitution of O., Ameyama, M. Purification and gluconate oxidase. J Biochem 1979, 86, characterization of 2-keto- D -gluconate 249–256. dehydrogenase from Gluconobacter Matsushita, K., Yamada, M., melanogenus. Agric Biol Chem 1981, 45, Shinagawa, E., Adachi, O., Ameyama, 1079–1085. M. Function of ubiquinone in the 100 Sonoyama, T., Tani, H., Matsuda, K., electron transport system of Kageyama, B., Tanimoto, M., Kobayashi, Pseudomonas aeruginosa grown K., Yagi, S., Kyotani, H., Mitsushima, K. aerobically. J Biochem 1980, 88, Production of 2-keto-L-gulonic acid from D -glucose by two-stage fermentation. 757–764. Appl Environ Microbiol 1982, 43, Shinagawa, E., Matsushita, K., Adachi, 1064–1069. O., Ameyama, M. Membrane-bound D -gluconate dehydrogenase of Serratia 101 Kondo, K., Ameyama, M. Carbohydrate marcescens. Purification and properties. metabolism of Acetobacter species. III. Agric Biol Chem 1978, 42, 2355–2361. Isolation and identification of D McIntire, W., Singer, T.P., Ameyama, lyxuronic acid produced from glucose by M., Adachi, O., Matsushita, K., Acetobacter melanogenum. Bull Agric Shinagawa, E. Identification of the Chem Soc Jpn 1958, 22, 380–386. covalently bound flavins of D -gluconate 102 Adachi, O., Tayama, K., Shinagawa, E., dehydrogenase from Pseudomonas Matsushita, K., Ameyama, M. aeruginosa and Pseudomonas fluorescens Purification and characterization of and of 2-keto- D -gluconate membrane-bound aldehyde dehydrogenase from Gluconobacter dehydrogenase from Gluconobacter suboxydans. Agric Biol Chem 1980, 44, melanogenus. Biochem J 1985, 231, 503–515. 651–654.

References 103 Ameyama, M., Osada, K., Shinagawa, E., Matsushita, K., Adachi, O. Purification and characterization of aldehyde dehydrogenase of Acetobacter aceti. Agric Biol Chem 1981, 45, 1889–1890. 104 Muraoka, H., Watanabe, Y., Ogasawara, N., Takahashi, H. Purification and properties of coenzyme-independent aldehyde dehydrogenase from the membrane fraction of Acetobacter aceti. J Ferment Technol 1981, 59, 247–255. 105 Fukaya, M., Tayama, K., Okumura, H., Kawamura, Y., Beppu, T. Purification and characterization of membranebound aldehyde dehydrogenase from Acetobacter polyoxogenes sp. nov. Appl Microbiol Biotechnol 1989, 32, 176–180. 106 Hommel, R., Kleber, H.P. Properties of the quinoprotein aldehyde dehydrogenase from “Acetobacter rancens”. J Gen Microbiol 1990, 136, 1705–1711. 107 Takemura, H., Tsuchida, T., Yoshinaga, F., Matsushita, K., Adachi, O. Prosthetic group of aldehyde dehydrogenase in acetic acid bacteria not pyrroloquinoline quinone. Biosci Biotechnol Biochem 1994, 58, 2082–2083. 108 Turner, C., Vela, C., Thony-Meyer, L., Meile, L., Teuber, M. Biochemical and genetic characterization of the acetaldehyde dehydrogenase complex from Acetobacter europaeus. Arch Microbiol 1997, 168, 81–91.

109 Lehmann, M., Tshisuaka, B., Fetzner, S., Lingens, F. Molecular cloning of the isoquinoline 1-oxidoreductase genes from Pseudomonas diminuta 7, structural analysis of IorA and IorB, and sequence comparisons with other molybdenumcontaining hydroxylases. J Biol Chem 1995, 270, 14420–14429. 110 Adachi, O., Shinagawa, E., Matsushita, K., Ameyama, M. Preparation of cells and cytoplasmic membrane of acetic acid bacteria which exclusively contain quinoprotein aldehyde dehydrogenase. Agric Biol Chem 1988, 52, 2083–2084. 111 Zahn, J.A., Bergman, D.J., Boyd, J.M., Kunz, R.C., Dispirito, A.A. Membraneassocciated quinoprotein formaldehyde dehydrogenase from Methylococcus capsulatus Bath. J Bacteriol 2001, 183, 6832–6840. 112 Klein, C.R., Kesseler, F.P., Perrei, C., Frank, J., Duine, J.A., Schwartz, A.C. A novel dye-linked formaldehyde dehydrogenase with some properties indicating the presence of a proteinbound redox-active quinone cofactor. Biochem J 1994, 301, 289–295. 113 Colby, J., Dalton, H., Whittenbury, R. Biological and biochemical aspects of microbial growth on C1 compounds. Annu Rev Microbiol 1979, 33, 481–517. 114 Shinagawa, E., Toyama, H., Matsushita, K., Tuitemwong, P., Theeragool, G., Adachi, O. A novel type of formaldehydeoxidizing enzyme from the membrane of Acetobacter sp. SKU 14. Biosci Biotechnol Biochem 2006, 70, 850–857.

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2 Catalytic Applications of Laccase Feng Xu, Ture Damhus, Steffen Danielsen, and Lars Henrik Østergaard

Summary

Laccases (EC 1.10.3.2) form a family of multi-copper oxidases capable of catalyzing the oxidation of many redox-active substances (particularly phenols) with the concomitant reduction of O2 to water. The enzymes, detected in various bacteria, fungi, plants, and insects, are postulated to play an important role in many cellular or microbial activities such as biosynthesis of plant cell wall, phytopathogenesis, degradation or humification of wood material, and pigmentation of bacterial/fungal spore. Extensive characterization of laccase has been carried out in the past decades. Since several comprehensive reviews have been written [1–13], we will focus in this chapter on the recent progress (mostly since 2000) in applying laccases as industrial catalysts.

2.1 Properties of Classical Laccase 2.1.1 Structure

Based on their structural and mechanistic properties, laccases belong to the multicopper oxidase family, along with bilirubin oxidase, ascorbate oxidase, ceruplasmin, and other homologs including those found in bacterial spore coat and copper resistance/efflux machinery [14, 15]. Numerous laccases have been cloned or are known for genome sequences. The conserved 10 histidines (His) and one cysteine (Cys) residues enable these proteins to possess four catalytic copper atoms, conventionally classified into three types according to the copper’s coordination and spectroscopic properties. The extracellular fungal laccase is archetypical. In general, a fungal laccase has a molecular mass of ∼60–80 kDa and an isoelectric point pI of ∼4–7, depending on glycosylation [1]. Of the four copper ions, the type 1 Cu (T1, ligated by at least one Cys and two His) has a strong electronic absorption around 600 nm (its

44

2 Catalytic Applications of Laccase Table 2.1 General molecular properties of typical laccases.

Source

kDa

pI

Glycosylation

Cu

A 280/A 600

Eº (T1) (V)

Fungal Bacterial

∼60–80 ∼20–80

∼4–7 ∼6–8

∼5–30% None

4/subunit 4/subunit

∼20 ∼10–20

∼0.4–0.8 ∼0.5–0.9

relative intensity to that at 280 nm, A 280/A 600, may be used to gauge the purity of the resting laccase) and a characteristic EPR signal. The type 2 Cu (T2, ligated by two His) also has a characteristic EPR signal, and the strongly coupled type 3 Cu pair (T3, each ligated by three His) has a weak UV absorption around 330 nm and is EPR active only when perturbed by strong anion binding [13]. Being an oxidoreductase, laccase’s Cu sites have defined redox potentials (Eº). For the T1 and T3 Cu, some laccases have a “low” Eº of ∼0.4–0.5 V (versus the normal hydrogen electrode), while others have a “high” Eº of ∼0.7–0.8 V [8]. However, the Eº of the T2 Cu appears to be ∼0.4 V for both the low- and high-Eº laccase groups [16]. Table 2.1 summarizes the general molecular properties of laccases that are often relevant for their applications. High-resolution crystallographic structures of a few fungal and bacterial laccases, including those of certain enzyme–substrate complexes, have recently been determined [17–22]. In general, three distinct domains in the folding of the backbone polypeptide chain are found. Combined with protein engineering work [23–27], the structural insight gained could enable us to further understand and engineer laccases.

2.1.2 Enzymology

It is believed that laccase catalysis involves (a) a reduction of T1 Cu by the reducing substrate, (b) an internal electron transfer from the T1 Cu to the T2/T3 trinuclear Cu cluster, and (c) a reduction of O2 to water at the T2/T3 Cu cluster [6, 11, 13]. The T1 and T3 Cu are linked mainly by a His-Cys-His tripeptide, whose Cys ligates the T1 Cu and whose His ligate two T3 Cu, and the T2/T3 Cu are electronically coupled to form a trinuclear cluster. Having a confined access channel to and binding pocket at their T2/T3 Cu cluster, laccases strongly prefer O2 as their oxidizing substrate. Having a much more open and shallow pocket at their T1 Cu site, however, laccases have a low specificity towards their reducing substrates [17–22]. Substrates whose redox site resides on freely rotatable benzene, benzothiazoline, or moieties of similar dimensions may easily dock inside the T1 pocket. A wide range of redox-active metal complexes, anilines, thiols, and especially phenols can transfer electrons to laccases, given that their Eº is ∼1 V or less. A K M in the order of 0.1 mmol L −1

2.1 Properties of Classical Laccase

and a kcat in the order of 103 s−1 are often observed for a typical reducing substrate, and a K m in the order of 0.05 mmol L −1 and a kcat in the order of 102 s−1 are often observed for O2 [1, 8, 28]. For many reducing substrates, their reactivity tends to correlate with the difference between their Eº and that of laccases’ T1 Cu, suggesting an “outer sphere” type of electron-transfer mechanism in which the activation energy is regulated mainly by the thermodynamic driving force, the Eº difference (ΔEº) [29–34]. Compared with a low-Eº counterpart, a high-Eº laccase may not only possess a higher oxidation potency (to work on more recalcitrant substrates) but also oxidize a substrate faster, making such an enzyme more attractive as an industrial catalyst. Laccases are often able to oxidize substrates with an Eº exceeding that of their T1 Cu, because the apparent endothermic oxidation half-reaction may be compensated by the vastly exothermic O2 to H2O reduction half-reaction, yielding an overall negative Gibbs’ free energy change. However, such energetics would diminish at alkaline pH, when the pH-sensitive Eº of O2/H2O is lowered close to or below that of laccases. For instance, the Eº of O2/H2O is ∼1.0, 0.8, and 0.6 V at pH 4, 7, and 10, respectively. In the range of pH 4–10, the Eº (T1) of Trametes villosa and Myceliophthora thermophila laccase is about 0.8 and 0.5 V, respectively [32]. Thus, thermodynamically, T. villosa laccase would become inactive above pH ∼7 to oxidize a substrate with an Eº > 0.8 V, and M. thermophila laccase would become inactive above pH ∼11 to oxidize a substrate with an Eº > 0.5 V. Sometimes, an initial endothermic electron transfer from a high Eº substrate to laccase may also be compensated by coupled chemical reactions (for example, deprotonation of an N—OH cation radical) [35]. In general, a bell-shaped pH-activity profi le (with optimal pH (pHopt) at ∼5–7) is observed for phenols, anilines, or other substrates whose oxidation by laccases is accompanied by H + dissociation. Because of the oxidative H + release, the Eº of these substrates decreases as pH increases. The subsequent increase of the ΔEº with laccase enhances the enzymatic oxidation, contributing to the ascending part of the pH profi le. At higher pH, however, the laccase inhibition by OH− becomes more pronounced, contributing to the eventual descent of the pH profi le [29, 32, 36]. For 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), hexacyanidoferrate(4), or other substrates whose oxidation by laccase is not accompanied by H + dissociation, a monotonic pH-activity profi le is often observed within pH 4–9, attributable to the relative insensitivity of their Eº to pH. Protonation–deprotonation of the substrates and/or laccase might also affect the pH profi le. Most fungal laccases are mesophilic, with optimal temperature (Topt) at ∼60 ºC. These laccases could quickly be inactivated at temperature above ∼50–60 ºC [1–3, 5–10, 12]. The thermal instability might be caused by protein unfolding or Cu loss. However, a laccase from the thermophilic Chaetomium thermophilum has a Topt of ∼70 ºC [37]. Laccase can be inhibited by various reagents, including halides, sulfanyl groups, and cationic quaternary ammonium surfactants [8]. Small “hard” anions such as

45

46

2 Catalytic Applications of Laccase Table 2.2 General enzymological properties of typical laccases.

Source

KM (mmol L-1)

kcat (s -1)

KM (O2) (mmol L-1)

pHopt

Topt (ºC)

Fungal Bacterial

∼10 −2–10 0 ∼10 −1–10 0

∼102–10 4 ∼101–102

∼10 −1 ?

∼5–7 up to 9

∼60 ∼80

F−, OH−, CN−, and N3− can tightly bind to the T2 Cu, interrupting the internal electron transfer and/or O2 activation. Thiols and sulfur-liking metal ions such as Hg2+ can cleave the T1 Cu-Cys ligation. Reductants may also leach Cu out from laccases. Table 2.2 summarizes some of the enzymological properties of laccase often relevant to its applications. 2.1.3 As Industrial Catalysts 2.1.3.1 Advantages Like many applied enzymes, a laccase catalyst should operate under ambient conditions, thereby save energy, and is biodegradable. Compared with peroxidases and other oxidoreductases, laccases may potentially have additional advantages as industrial catalysts: 1. Laccases use readily available, inexpensive, and safe O2 as co-substrate. 2. Laccases can oxidize a wide range of molecules (as their reducing substrates). These include phenols, anilines, thiols, N-hydroxyls, N-oxides, N-oximes, phenazines, phenoxazines, phenothiazines, transition metal complexes, and so forth (Table 2.3). Many of the compounds occur naturally and/or can be important industrial precursors, products, or byproducts. Their transformability by laccases might allow various viable applications of these enzymes. 3. Laccases can be produced industrially by fermentation. Many fungal laccases can be secreted by hosts engineered to remove undesirable contaminants and/or toxins, making their recovery and formulation relatively straightforward. Peroxidases and some other oxidoreductases often require complex cofactors (for example, heme and flavin), whose biosynthesis may severely limit the overall expression of the holoenzyme. Supplementing these cofactors exogenously could be prohibitively expensive. In contrast, laccase employs Cu as cofactor, which can be readily supplied by simple, inexpensive Cu salts.

2.1 Properties of Classical Laccase Table 2.3 Types and examples of molecules directly oxidizable by laccases.

Phenols, anilines, benzenethiols Methyl syringate

HO H3CO

OCH3

COOMe

3-Hydroxyanthranilic acid

NH2 OH

HOOC

1,2-Phenylenediamine

H2N NH2

Benzene-1,2-dithiol

SH SH

N-Hydroxyls/oxides/oximes N-Hydroxybenzotriazole

OH

N

N N

N-Hydroxyacetanilide

OH N COCH3

Violuric acid

OH

O

N

HN O

O

N H

N-Hydroxyphthalimide

O N O

TEMPO N

+

O-

OH

47

48

2 Catalytic Applications of Laccase Table 2.3 Continued

Phenoxazines, phenothiazines Phenoxazine-10-propionic acid

O

N COOH

Phenothiazine-10-propionic acid

S

N COOH

Other redox-active organic compounds ABTS

H3C HO3S

N

S N

N

N

S

SO 3H

CH3

Redox-active metal complexes K4Fe(CN) 6, Fe(C5H5) 2, Fe(bipyridyl)3Cl2, K4Mo(CN) 8, Mn(oxalate)

2.1.3.2 Shortcomings For general oxidation applications, laccases may be inferior to peroxidases in terms of oxidation potency, because their Eº (∼0.4–0.8 V) is relatively low compared with that of heme peroxidases (∼1 V). Laccases may also be inferior to oxygenases in terms of stereo (particularly enantio)selectivity when applied for synthesis, because they oxidize substrates by abstracting electrons (to yield radical products), rather than forming covalent intermediates. In general fungal laccases are destabilized at temperatures above ∼50–60 ºC (even though the enzyme could withstand alkaline pH up to ∼10 at room temperature). Typical laccases are often inactive toward insoluble substances or oligo/polymerized soluble substrates [38], although this may be overcome by using mediators (see Section 2.2.1.6).

2.2 Applications of Laccase for Industrial Oxidation Processes

The ability of laccases to catalyze the oxidation (by O2 ) of various substances of industrial and other interest has driven intensive research efforts to exploit and develop these enzymes as industrial catalysts. Based on the aims, the reported application trials could be divided mainly into degradation (of waste, pollutants,


and so forth), transformation (of raw materials into functional materials), and sensing, covering industrial-technical, specialty chemical synthesis, environmental, food, medicinal and personal care fields. Laccases are exploited to replace hazardous/expensive chemicals, save on energy/resources consumption, create novel functionalities, and reduce detrimental impacts on the environment. 2.2.1 Laboratory-Level Trials 2.2.1.1 Delignification As a major enzyme type secreted by white-rot fungi, laccases have been extensively studied for delignifying pulp. Although the conventional chlorine- or oxygen-based chemical oxidants are very effective, they can cause serious problems in byproducts disposal or cellulose fiber strength loss. Laccases might be applied to oxidatively degrade lignin, so that the use of chemical oxidants could be reduced [39]. Various laccases are able to degrade both natural and synthetic lignin [7, 40–45]. Most of the studies in the field have been carried out on monomeric or oligomeric model compounds (for recent reviews, see [7, 29, 44, 47, 48]). Experimental data suggest that in general, laccase first oxidizes accessible phenolic moieties in lignin to generate phenoxy radicals. From these radicals, quinone formation, crosslinking, or other reactions may take place. In the presence of a redox mediator, the non-phenolic moieties in lignin (particularly the benzylic Cα site) may also be oxidized, leading to other reactions such as peroxygenation, demethoxylation, Cα-Cβ cleavage, Cβ-O homolysis, or aromatic ring opening [39, 49–51]. Laccases alone tend to polymerize lignin, by generating cross-linking, Ocentered phenoxy radical (Ph-O•). With a mediator, however, laccase catalysis tends to lead to lignin depolymerization, likely due to the action of bond-cleaving C radicals [29, 52, 53]. Delignification of pulp by laccase may be demonstrated by kappa number reduction or improvement for the downstream chemical bleaching. Mediated laccase systems, particularly those based on N-hydroxy mediators, have made significant strides for delignification application. Satisfactory pulp bleaching could be achieved (for recent reviews, see [7, 47, 48, 52]). However, major hurdles still exist for future commercialization of the technology. Currently, the mediator has to be loaded at ∼0.1–1% w/w based on pulp, a level too expensive for the industry [46, 54]. The high loading may also pose disposal problems for spent mediators, whose transformation during pulp bleaching needs further study for potential environmental impact. It seems that some white-rot fungi employ highly efficient, natural laccase mediators in wood degradation. It is highly desirable to discover novel, natural, or artificial mediators having high reactivity as well as high turnover stability, so that the mediator dosage for laccase-based pulp delignification may be reduced by several orders of magnitude to make this technology economically viable [44, 55, 55a].

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2 Catalytic Applications of Laccase

2.2.1.2 Dye and Colorant Bleaching Dyes and colorants are diverse molecules with many unsaturated bonds and/or chromogenic groups whose interaction give rise to different colors. They are often prone to laccase oxidation, which in some cases lead to decolorization. Thus laccases have potential for dyeing, waste treatment, and paper recycling applications [1, 2, 5, 6, 8, 9, 48, 56, 56a]. Research on the ability of laccases to oxidize acid, anthraquinoid, azo, indigoid, heterocyclic, phthalocyanine, stilbene, triarylmethane, and other types of dye continues [42, 57–62]. Using mediators may extend laccase oxidation to more recalcitrant dyes, with efficacy depending on the reactivity and stability of selected mediators [58, 63–65]. Immobilizing a laccase may enhance its stability, resistance to inhibitors, or recyclability [42, 61]. In addition to being decolorized or degraded, dyes may be polymerized or attached to other substances (for example, humic acid), allowing easy physical removal. Degrading the dyes in dyehouse effluent by laccase oxidation could significantly reduce environment pollution, enhance water reuse, or prevent undesirable crossdyeing. Bleaching the dyes in recycled papers by laccase could reduce the overall cost. Applying laccase to the finishing of dyed denim has been commercialized, in which the enzyme is used to bleach the indigo dye to achieve a desirable appearance of the final product (see Section 2.2.2.2). 2.2.1.3 Bioremediation Industrial wastes, agricultural (biocides) residues, and municipal discharges leave numerous pollutants, xenobiotics, and other hazardous chemicals in the environment. Active research is conducted to evaluate the ability of laccases to oxidatively detoxify or degrade these compounds, such as bisphenol A or 2,4dichlorophenol (endocrine-disrupting or estrogenic), cyprodinil (fungicide), chlorinated phenol or α-naphthanol (pesticide leftover), and various aromatics [65a, 66–72]. The laccase catalysis could result in either degradation, polymerization, or immobilization of the target molecules [1, 2, 5, 6, 9, 48]. For instance, laccases may dehalogenate or oligomerize chlorophenols to reduce their toxicity. Laccases may also polymerize xenobiotics as such or copolymerize them with other substances (for example, humic acid) to facilitate molecular mass-based removals such as adsorption, sedimentation, or filtration. Laccases have also been studied for degrading sulfur-containing coal substances, for potential applications in coal biodesulfurization/upgrading, bioremediation around coal mines, and emission reduction of acid rain-causing agents from power plants [4, 8]. To commercialize such bioremediations, many technical hurdles need to be overcome. Among them, immobilizing laccase on a solid or gel carrier seems to be a prerequisite, considering the needs of using the enzyme in numerous cycles (to reduce cost) and/or in heterogeneous multi-phase systems (to enhance the contact between the water-soluble enzyme and insoluble substrates).


2.2.1.4 Other Degradation Applications Chemical oxidants such as hypochlorite are widely used in detergent/washing solution to bleach/degrade stains from fabric, dish, or other surfaces. It is of interest to replace the chemicals with enzyme (including laccase)-based oxidants, although little progress has been made so far [73, 74]. Laccase may also be used to whiten cotton fiber, by oxidatively bleaching various native or acquired chromophores [75]. Many body, domestic, and industrial odors are caused by sulfides, thiols, ammonia, amines, short chain fatty acids, or other volatile organic compounds. Being able to oxidize various thiols and other S compounds, laccases have been studied for deodorant applications including those that target pulping plant odor and halitosis [76–79]. Rather than simply masking the malodor with fragrances as conventional deodorants do, a laccase system could degrade the offensive molecules, or even kill the microbes that generate them. Laccase may be used for crop/plant protection, by oxidatively degrading toxins secreted by invading microbes, such as the cercosporin from Cercospora [80]. Browning, hazing, and turbidity during the processing or storage of clear fruit juice, beer, and wine can be major problems for the industry. It is believed that phenolic compounds, such as epicatechin, ferulic acid, and coumaric acid, are involved. Conventional fining agents (for example, gelatin and bentonite) used to adsorb these phenolics usually have low specificity, may affect color/aroma, or can pose disposal problems. Laccases may oxidize and polymerize the problematic phenolics, making their removal by silicate fining more easily, so that the appearance, flavor, aroma, taste, or stability in fruit juice or fermented alcohol beverages could be improved [8, 81, 81a, 82]. The consumption of O2 with formation of water makes laccases attractive in some cases as a deoxygenating agents for food preservation [83]. The progress made recently in applying laccase to degrade recalcitrant chemicals, such as polycyclic aromatic hydrocarbons (PAHs) and chemical warfare agents, will be discussed in Section 2.3.2.2. 2.2.1.5 Functional Biotransformation Modified lignocellulosics can have many uses as novel composites or other functionalized materials. Laccases could transform low-value lignocellulosics in three ways: (a) by directly oxidizing and cross-linking wood particles or pulp, (b) by grafting particular molecules (such as aromatics, carboxylic acids, isocyanates, and acrylamide) onto wood particle or pulp surfaces, and (c) by transforming isolated lignin into radical-rich adhesives [1, 5, 6, 8, 84]. Such applications of laccase could not only replace toxic or expensive chemical adhesives or processing aids/additives, but also upgrade low-value forestry byproducts. Recent progress has been made in activating the fiber surface of thermomechanical pulp (by generating radicals from the residual lignin via the mediation of dissolved/colloidal wood substances) [85, 86], enhancing paper tensile strength (via intrasheet cross-linkage of generated phenoxy radicals) [87, 88], grafting acrylamide onto

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pulp for novel composite/plastic [89, 90], and making fiber-board without ureaformaldehyde adhesive [91]. To commercialize such enzymatic applications, further optimization is needed so that they can be competitive against the inexpensive chemical counterparts. The radical-generating ability of laccases seems well suited for polymerization reactions. For example, they may polymerize phenols and anilines into resins, replacing conventional chemical couplers such as formaldehyde [91a, 91b]. Conductive polyaniline polymers, useful for various electrical or electronic applications, may also be made using a laccase [92]. Urushiols, which are catechol derivatives with alkene/allylic side-chains, often found abundantly in tree sap, may be oxidatively polymerized by laccases into artificial urushi (Japanese lacquer) or other bio-based products [93]. Being able to oxidize urushiol could also make laccases effective as topical agents against poison ivy dermatitis, caused mainly by urushiol [8]. Laccases may be applied to oxidatively transform and subsequently couple dye precursors in situ to cellulose in textile, keratin in hair, or collagen in hides [8, 94–96]. The process could improve the dyeing/tanning or reduce the cost (by using less expensive precursors). Particularly, laccase-based hair dyeing or waving processes using less harsh chemicals and/or milder conditions appears attractive [4, 8, 97]. Laccase-catalyzed modification of humic acid may have potential uses for regulating the dynamics of soil organic matter [98], in addition to pollutant-removal. Some laccase substrates, such as phenols and thiols, are important components of various foods. Their modification by laccase may lead to new functionality, quality improvement, or cost reduction [81, 99]. For instance, laccases may oxidize and/or cross-link the tyrosyl groups in myofibril proteins, leading to rheological changes of meat products [100, 101]. The ferulic acid units in arabinoxylans, pentosans, or pectins may be oxidized by laccase, leading to the gelling of cereal foods [102, 103]. Whey proteins and α-lactalbumin may be cross-linked by laccase in the presence of chlorogenic acid as a mediator. Gluten or other wheat components may also be affected by laccase in baking applications [104]. Many intermediates or products generated by laccases could have a biocidal effect in disinfection or personal-care fields. For instance, laccase can oxidize iodide to iodine, a widely used disinfectant [105], or generate microbe-killing phenoxy or N-oxy radicals. Such a laccase-precursor disinfection system may be quickly turned on by hydrating/mixing, and be used for in situ sterilization. Laccase can also transform 3-hydroxyanthranilic acid (Table 2.3) to cinnabarinic acid, an antibacterial agent effective on Gram-positive streptococci [106]. The enzyme has been studied for synthesizing various bioactive compounds, such as dimeric curcuphenol, estradiol, resveratrol, imidazole-based compounds, catechin-hydroquinone adducts, or dihydrocaffeic acid derivatives [99, 107–111]. A recent study applied such laccase biocatalysis to crop/plant disease control [112]. Laccase may be used to synthesize several complex medicinal agents, including vinblastine, penicillin X dimer, cephalosporin, triazolo(benzo)cycloalkyl thiadi-


azines, and dimerized vindoline [8]. There is potential to apply laccase against biological and/or chemical warfares (see Section 2.3.2.1). Recently exploited laccase applications in organic synthesis will be discussed in Section 2.3.2.4. 2.2.1.6 Biosensing Many substrates of laccases are either chromophoric, chromogenic, fluorogenic, chemiluminescent, or electroactive. They may be converted to products suitable (either directly or indirectly via coupled chemical or enzymatic reactions) for optic, electric, or other physical measurements. Thus laccase may be applied as biosensor or bioreporter. Laccase catalysis, coupled with various physical transducers, could be useful for detecting O2 and a wide variety of reducing substances, especially phenols or anilines [8]. One type of such biosensor may monitor electric signal changes from a modified electrode under the electrocatalysis of a laccase. Another type may detect the photometric change resulted from the oxidation of a chromophoric/ chromogenic substrate. Recent studies include the amperometric detection on laccase-modified electrodes of morphine, phenols, and flavonoids for doping detection, waste monitoring, and beer quality control, respectively [113–116, 116a]. Laccase catalysis may also be used to assay other enzymes. In these assays, either the enzyme of interest catalyzes the production of a compound whose subsequent transformation by laccase generates a detectable physical change, or a product from the laccase catalysis (whose production is accompanied by a detectable physical change) is quenched by the activity of the enzyme of interest. The strategy has been applied to assay various hydrolases, transferases, and oxidoreductases [8]. Laccase covalently conjugated to bio-binding molecules (for example, antibodies, antigens, DNA, RNA, biotin, or streptavidin) may be used as a reporter for immunochemical, histochemical, cytochemical, or nucleic acid detection assays [8]. In these assays, the binding moiety binds to the target, and the laccase’s reaction signals the binding event. Recent progress includes a sandwiched immunosensor and a virtually substrate-free insulin sensor [117, 118]. 2.2.1.7 Desirable Application Modes To become viable industrial catalysts, laccases need to be subjected to various treatments in order to make them robust, recyclable, or heterogeneous. One of the most studied treatments is immobilization, achieved by either chemical linkage or physical adsorption/entrapment in various carriers [119]. Covalent coupling to a solid carrier (for example, celite, alumina, or nylon [7, 61, 120]) may allow multiple uses of the laccase (thus saving in enzyme cost or water usage) or increase its resilience against pH or thermal inactivation, although the immobilization may lead to a significant activity loss of the original enzyme. However, many targeted molecules or their products (particularly dyes) may also adsorb onto the carrier, resulting in either inactivated catalyst, inaccessible substrate, or inseparable product [57, 119]. Encapsulation in hydrated gels, for example Cu-

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alginate [59], may better preserve laccase, although the hindered permeability or unproductive adsorption of a substrate or product may pose problems. Entrapment in micelles or immobilization on solid carriers may allow laccases to be applied in non-aqueous, novel solvents (organic solvents, ionic liquids, or supercritical CO2 ) or multiphasic systems. Other kinds of modification have also been studied on laccases. Cross-linking laccases into enzyme crystals could enhance their stability [121]. Wrapping laccases (via covalent surface linkage) with amphiphilic polymers [122], dendrimers [123], PEG [124, 125], or a dioleyl glucono glutamate surfactant [126] could protect them from proteolysis, facilitate their incorporation into micelles, solubilize them in organic solvents (particularly the non-aliphatic, reverse micelle-disrupting ones), or enhance their accessibility toward hydrophobic substrates. Non-ionic, cationic, and anionic polymers (including surfactants) tend to exhibit different effects on laccase catalysis, attributable to their different function in binding product or laccase [127–129, 129a]. Combination of laccase catalysis with other physical or chemical methods, such as electrochemistry and ultrasonics, has been studied [130, 131]. For applications that target large, insoluble, or non-substrate substances, small redox-active mediators have been found very useful in enhancing/extending laccase catalysis. This is exemplified by the laccase-mediator systems for pulp delignification (in which the targeted lignin is insoluble, hardly accessible, and mostly inactive towards laccase direct action) and for dyed fabric finishing (in which the targeted dye is adsorbed onto insoluble textile, making it mostly inaccessible and inactive for laccase). Numerous molecules have been studied as potential laccase mediators. Currently promising mediators include some types of phenol/aniline [7, 42, 132, 133], N-hydroxy/oxide/oximes [30, 31, 42, 54, 134– 136], phenazine/phenoxazine/phenothiazine [31, 42, 46, 134], and redox-active organics or metal complexes [29, 46, 64, 134, 137–139]. Mediated laccase catalysis has potential for the degradation or modification of lignin [39, 42, 47, 49, 51, 63, 84], delignification of pulp or bleaching of recycled paper [54, 88], degradation of xenobiotics and other pollutants [65a, 69, 71], bleaching of dyes [60, 64], and synthesis of fi ne or bioactive chemicals [108, 134, 141]. Table 2.4 shows some of the types of targets suitable for mediated laccase oxidation. In general, any reducing substrate of laccase having an Eº close to ∼1 V and a relatively stable oxidized state (or insignificant autooxidation) may serve as a potential mediator, capable of being oxidized/reoxidized by laccase and oxidizing target molecules (essentially shuttling oxidation equivalences between laccase and target). All the known mediators seem to be oxidized by laccase via “outersphere”-type electron transfer, whose kinetics is dominated by the ΔEº between the mediator and laccase (see Section 2.1.2). Based on the proposed mechanisms for oxidizing the target, known laccase mediators may be divided into three main groups [35, 142]. The first group, including ABTS, phenols, and phenothiazines (Table 2.3), seems to oxidize a target by electron transfer. For instance, laccasegenerated ABTS radical and doubly oxidized form may extract electrons from a phenol and non-phenol target, respectively, with a rate/efficiency proportional to


55

Table 2.4 Potential targets for mediated laccase biooxidation.

Representative mediator

Representative target (targeted bond/atom in bold)

ABTS

R'

O X

R'O

P

S

R

SR

OR"

Phosphorothioate N-Hydroxybenzotriazole

Chlorobenzene

R

C

H

C

Toluene thiophene R

H

C

H H

H H R

C

R'

Benzylic alcohol/amine TEMPO

O

H

Phenylpropene

Cyclohexene

Linoleic acid

O H

O OH OH

OH OH

Glucose

Cyclohexylmethanol

the ΔEº between the mediator and target [29, 140, 143, 144, 144a] or attributable to the Hammett correlation with regards to the electronic property of the target’s substituents [64]. The second group, including N-hydroxybenzotriazole (HBT), N-hydroxyacetanilide, and other N—OH compounds, seems to oxidize a target by H-abstraction. For instance, the laccase-generated N—OH + • cation radical in HBT may deprotonate into an N—O• radical, which then abstracts an H from a benzylic target to form a benzylic C• radical, with an efficiency dependent on the bond dissociation energy (BDE) difference [35, 135, 143–145]. The third group, represented by the radical 2,2,6,6-tetramethylpiperidin-1-yloxyl (TEMPO), seems to employ yet another mechanism (see Section 2.3.1.2). It is the second and third groups of mediators that have significantly extended laccase catalysis to nonsubstrate substances. 2.2.2 Commercialized Applications

Built on many years of basic and applied research, laccases started being commercialized as industrial catalysts in the late 1990s. Besides use as analytic or diagnostic agents, there are two established industrial applications for laccase in the fields of wine bottling and textiles [4, 74], and two nascent applications in the fields of waste treatment and cardboard packaging materials.

R'

56


2.2.2.1 Preventing Taint in Cork Stoppers It is a well-known problem that cork stoppers in bottled wine may, apparently more or less at random, impart an unpleasant flavor to the wine with an astringent mouth feel and/or a musty taste. Bottles with such a taint must be considered as spoiled. As far as the astringency is concerned, the culprit seems to be the phenolic compounds (including chlorinated phenolics) in the cork. One particular compound, 2,4,6-trichloroanisole, with an extremely penetrating taste, is suspected of giving the musty character [146, 147]. These components may be jointly pacified by enzyme-catalyzed polymerization of the phenolics in a process where the corks are washed with a laccase solution, thereby presumably either washing out or copolymerizing/entrapping the anisole as well [148]. Cork surfaces may also be rendered more hydrophobic by the enzymatic polymerization, resulting in better water repulsion and/or reduced leakage of cork substances into wine [149]. A commercial laccase product, Suberase® from Novozymes, has been developed and marketed for effecting such a treatment. 2.2.2.2 Denim Bleaching Laccase-catalyzed textile dye-bleaching has found its use in finishing indigo-dyed cotton fabric, denim [150–155, 155a]. Commercial laccase products from Novozymes, first Denilite® and later Denilite II®, contain a fungal laccase and a redox mediator [74, 150]. With the help of the mediator, the laccase can effectively catalyze the oxidation of the (insoluble) blue indigo into the essentially colorless isatin. The traditional chlorine-based bleaching, applied as the fi nal step in finishing of indigo-dyed denim when the fashion requires a bleached look, is replaced by a much more environmentally acceptable method, whose benefits include better processing control, reduced water usage, and less pollution. This application is an interesting case from the point of view of enzyme technology development. Mediated oxidoreductase (peroxidase or laccase) systems have been actively pursued for laundry detergent applications, particularly for “dye-transfer inhibition” during washing (or bleaching of fugitive textile dyes in the wash liquor) [56]. The research encountered a major difficulty in how to bleach the unwanted, desorbed dye while preventing fading of the dyed garments. Although a mediated laccase system for household laundry has yet to be developed due to this difficulty, the research was redirected towards an application where bleaching of dyed garments is desired, leading to the successful laccase product for denim bleaching in the textile industry. 2.2.2.3 Paper Mill Effluent Treatment and Cardboard Strengthening As of recently, a laccase product (Novozyme 51003) is applied for treatment of paper mill effluent. The effect of adding the laccase is to reduce the color and biochemical or chemical oxygen demand in the effluent and to improve the biological step in the sewage treatment plant, all presumably due to laccase-catalyzed polymerization of lignin components in the effluent (Henrik Lund at Novozymes A/S, personal communication, June 2006).

2.3 More Recent Developments

The laccase product is also used recently for strengthening packaging materials made from old corrugated cardboard, probably by affecting the cross-linking of cellulosic fibers via reaction with lignin components present on the fiber surfaces (Henrik Lund at Novozymes A/S, personal communication, June 2006). 2.2.2.4 Major Hurdles to Further Development from Laboratory Trials To further develop a laccase catalytic system (from its promising laboratory-level study) may involve many scientific, technical, engineering, economic, environmental, or other hurdles. One major hurdle could be the cost of laccase and/or mediator. To reduce the cost of laccase, higher yield for fermentatively produced laccase or novel laccase productions (see Section 2.4.2) is needed. To reduce the cost of a mediator, controlling its unproductive side-reactions is necessary. For instance, the prohibitively high dosage of the currently best N—OH mediators needed for delignification (∼0.1–1% w/w of pulp) is mainly caused by the significant dehydroxylation or deoxygenation side-reaction (>N—OH or >N—O• → >N—H) and the consequent low turnover of the mediator [39, 42, 47, 49, 51, 54, 63, 84]. Another major hurdle could be the inactivation of laccase or mediator in the highly heterogeneous or variable application environment, whose abundant inhibitors/reactants might not be present in defined laboratory systems [58]. The lack of comprehensive knowledge with regard to the environmental impact of the mediator (particularly when it is synthetic) and/or its derivatives may also hinder the application [42, 132].


Riding on the emerging wave of industrial biotechnology, the research on laccasebased catalysis is being expanded in both scope and depth. Discoveries of new laccase, mediator, and application leads have been made recently. 2.3.1 Novel Laccase Catalytic Systems 2.3.1.1 New Laccases Besides the archetypical laccases discussed in Section 2.1.1, various seemingly “unconventional” laccases or analogs have been reported. Some of the isolated laccases (with sequences highly homologous to typical multi-Cu oxidases) appear as “yellow” or “white” when purified, indicating the lack of a canonical “blue” T1 Cu site. The site might be modified by oxidized substrates, as proposed for the Pleurotus ostreatus or Panus tigrinus “yellow” laccase [157, 158], or contain other metal ions (for example, Zn2+ or Fe3+ ), as proposed for the P. ostreatus “white” laccase [159]. Often, a “yellow” or “white” laccase may oxidize non-phenolic compounds in the absence of mediator, or exhibit a more desirable pH or thermal profi le than its “blue” counterpart.

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Recently reported laccase analogs, classified as such based on their catalyzed reactions, include a dimeric (43 kDa subunit) Tricholoma giganteum (mushroom) oxidase which also shows an anti HIV-1 reverse transcriptase activity [160], a dimeric (40 kDa subunit) Tellima grandiflora (plant) oxidase [161], and an 85 kDa Nephotettix cincticeps (insect) oxidase [162]. However, these oxidases have an Nterminus either unknown or not homologous to typical laccases. Thus more structural and enzymological information is needed to determine whether these oxidases belong to the laccase (or multi-Cu oxidases) family. The discovery of novel laccases is propelled by the ever-increasing numbers of sequenced genomes and the revealed genetic information. The huge amount of data has sparked development of tools for computational analysis to link sequence with biological function. These tools have naturally also been employed to study the classical laccases. More than 100 plant and fungal laccase sequences were thus analyzed using sequence alignment [15] to identify signature sequences for laccases. A phylogenetic analysis including more than 350 multi-Cu oxidases was reported in another study [14]. The largest group of enzymes in the latter study was indeed fungal laccases, which clearly separated in two clusters of sequences from basidiomycetes and sequences from ascomycetes. The sequences were selected for the study by the presence of four conserved Cu-oxidase consensus patterns and the study also included a number of bacterial protein sequences. These bacterial sequences clustered quite separately from all of the eukaryotic proteins. A couple of bacterial sequences were even excluded, as they could not be aligned without loss of resolution in the phylogenetic analysis. The example serves to illustrate the difficulty in classification of bacterial laccase sequences that have emerged amongst the multicopper oxidases in the last decade. The Pfam database is a collection of protein families and domains and it contains multiple protein alignments and profi le-HMMs of these [163]. In Pfam, the multicopper oxidases constitute a clan with seven Pfam members. The classical fungal laccases contain elements from no less than three members of this clan. Some of the new bacterial laccases differ from this archetype and have a completely different domain organization. The expansion of the laccase enzyme class with prokaryotic members clearly represents both a great opportunity and a challenge as well to the field [164, 165, 165a]. Bacterial laccase-like enzymes have been discovered in Azospirillum, Bacillus, Escherichia, Marinomonas, Pseudomonas, Streptomyces, Thermus, and Xanthomonas genera. Many bacterial laccase genes have a signal-like sequence, but the cellular localization and biological role of these enzymes is far from wellestablished. Some genes, such as the laccases from Streptomyces coelicolor and Thermus thermophilus, include a Tat-signal sequence, which labels the gene-product for translocation via the Tat-pathway [166, 167]. Indeed, S. coelicolor laccase activity has been detected in the media when overexpressed homologously. Laccase activity has in other instances been located to the surface of spores, such as those of Bacillus subtilis [17], where it is involved in production of pigmentation offering protection against UV radiation.


Comparison of amino acid sequences of bacterial laccases reveals a large degree of diversity. This variation is reflected in several molecular parameters of the laccases. The molecular weight spans the range from 32 kDa for S. coelicolor laccase [166] to approximately 65, 70, or 75 kDa for B. subtilis, S. lavendulae, or S. cyaneus laccase [168–170]. The higher levels of structural organization also differ. Some bacterial laccases, such as the polyphenol oxidase from Marinomonas mediterranea, are active as monomers [171], while others, such as the homotrimeric EpoA from S. griseus, are active with quaternary structure [172]. Based on sequence analysis, the small laccases from S. coelicolor and S. griseus likely lack the domain 2 (Pfam id: Cu-oxidase), which is present in all fungal laccases [17– 22]. The small laccases have a full complement of copper atoms and it is plausible that their multimeric organization somehow compensate for intradomain interactions otherwise present in laccases [166]. Consequently, the bacterial laccases may have (reducing) substrate clefts different from those in fungal laccases, resulting in different specificity, reactivity, stability, or other properties. However, the current molecular models do not provide sufficiently reliable information about such intermolecular interactions, and the 3D structures of these small laccases are yet to be elucidated. Some enzymatic properties of bacterial laccases are similar to those of fungal laccase (Tables 2.1 and 2.2). For instance, S. griseus laccase has a Topt of ∼40 ºC [172], T. thermophilus laccase has a pHopt of ∼5 when oxidizing phenols [167], and B. halodurans laccase has a bell-shaped and monotonic pH profi le for phenolic and non-phenolic substrates, respectively [173]. However, other properties could be quite different. The Eº (T1) differs significantly between the various bacterial laccases: S. coelicolor laccase has an Eº of only 0.5 V [166], while the Eº of M. mediterranea polyphenol oxidase is reported to be >0.9 V at pH 7 [171]. The laccases are all reported to oxidize a variety of phenolic and non-phenolic substrates, such as syringaldazine, dimethoxyphenol, ABTS and K4Fe(CN) 6. The laccase of S. griseus appears to be the exception as it oxidizes dimethoxyphenol but not syringaldazine [172]. A general trend of the bacterial laccases is their high degree of stability with the extremes being the laccase from T. thermophilus and S. coelicolor. The Topt for the T. thermophilus laccase is approximately 92 ºC and its half-life of thermal inactivation is more than 14 h at 80 ºC [167]. S. coelicolor laccase can retain activity after boiling and treatment with SDS [166]. Another striking feature of the bacterial laccases is their good catalytic activity under alkaline conditions. The activity of S. coelicolor laccase on dimethoxyphenol is reported to peak at pH 9.4 [166]. It appears that the catalytic efficiency above neutral pH is rate-limited by the noncatalytic interaction between the substrate and the T1 Cu site. It is therefore likely that bacterial laccases bind negatively charged substrates better than positively or uncharged molecules. Other bacterial or archaeon laccases also demonstrate “unusual” pH, thermal, or inactivation-resistance properties. B. subtilis endospore coat CotA laccase is active at high pH and temperature, likely related to the requirement of the spore survival under extreme conditions [17, 168]. The laccase

59

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has been applied in dye bleaching at pH 9 and 60 ºC, compared with ∼ pH 7 and 40 ºC suitable for fungal laccases [174]. S. lavendulae and S. cyaneus have a laccase quite stable at ∼70–75 ºC [169, 170]. A 56 kDa laccase from B. halodurans can be stimulated by NaCl, in contrast to the halide inhibition of fungal laccases [173]. In summary, the bacterial laccases constitute a highly diverse subgroup within the laccase enzyme class. The diversity is apparent at the amino acid sequence level, which is naturally reflected in various physicochemical parameters of the enzymes (for example, molecular weight, pI, Eº). The bacterial laccases differ from their fungal counterparts by enhanced temperature stability. Furthermore, they generally exhibit good activity above neutral pH. Their substrate specificity resembles that of the fungal laccases with a preference for aromatic phenolic substrates. Genome sequencing will undoubtedly lead to the discovery of even more bacterial laccases with properties different from the fungal laccases. The increase in diversity makes these laccases most interesting targets for protein engineering, either by site-directed mutagenesis [17] or gene shuffling (see Section 2.4.1). 2.3.1.2 New Mediators As discussed in Section 2.2.1.7, improved mediators are needed to further develop viable laccase catalytic systems. To do this, extensive work has been carried out to test derivatives of promising known mediators, such as ABTS, HBT, and phenothiazine-10-propionic acid, to systematically study their structure–function relationship [30, 31, 136, 144, 175]. Phenols with extended conjugation and/or steric structure causing low pK a have been studied to better balance their reactivity towards laccase/target with their stability at the oxidized form [137]. Pyrazolones have been investigated and found to be inferior to ABTS as potential laccase mediators [176]. Many redox-active metal complexes have been studied [29]. The Mn3+ /Mn2+ couple has been exploited, and the oxidation potency of Mn3+ depends on its chelator [177]. Polyoxometalates (like tungsto-vanado-silicates, Eº ∼0.74 V) have previously been used alone as robust redox catalysts for delignification. Their slow reoxidation by O2 is often rate-limiting, but may be accelerated under laccase catalysis [178, 179]. The most significant advance in the field comes from the discovery and characterization of TEMPO and its analogs as novel laccase mediators. The cyclic structure of TEMPO hosts a very stable N—O•, making the radical less active in H-abstraction than the N—O• of oxidized HBT or its derivatives. The BDE of the NO—H in N-hydroxy-2,2,6,6-tetramethylpiperidine is ∼290 kJ mol−1, which is lower than that in N-hydroxyacetanilide (∼330) and N-hydroxyphthalimide (∼370) [35, 180]. Since the BDE of a benzylic C—H is ∼340 kJ mol−1, abstracting this H by TEMPO would be unfavorably endothermic, in contrast to the H-abstraction by N-hydroxyphthalimide radical. Oxidized TEMPO, an oxoammonium (>N + ¨ O) species and a more potent oxidant, is well known for its potency/efficacy in oxidizing various molecules [180–182, 182a]. The oxoammonium has an Eº of ∼0.9 V, and may be readily made by an appropriate laccase [181, 183]. Thus TEMPO can serve as a mediator for laccase biocatalysis.


Unlike the electron-transfer or H-abstraction mechanisms for (laccase-) oxidized ABTS or N-hydroxy/oxide/oxime mediators (as discussed in Section 2.2.1.7), the mechanism of TEMPO oxoammonium in oxidizing target molecule is believed to be of a two-electron, “ionic oxidation” type. For an alcohol target, nucleophilic attack of the alcohol O on the N + of the oxoammonium forms a tetrahedral intermediate, followed by removal of an Hα in the alcohol (by TEMPO’s N—O− or a solute base under acidic or basic conditions, respectively), resulting in N-hydroxyl-2,2,6,6-tetramethylpiperidine and an aldehyde product. The hydroxylamine can be reoxidized by laccase to first TEMPO and then the oxoammonium form to start another round of alcohol oxidation [35, 144, 144a, 184]. Hence, TEMPO-mediated laccase catalysis may be applied on targets with ionization potential too high for a direct oxidation by laccase [2]. The involvement of a tetrahedral intermediate in the oxoammonium’s interaction with the target may also give the catalysis more stereoselective than other laccase oxidations. The TEMPO-mediated laccase biocatalysis has recently been exploited for delignification [39], dye bleaching [42], degradation of chlorinated aromatics [185], and phenolic oligomerization [38]. Despite TEMPO’s ability to mediate this unique type of laccase catalytic process, there is still a need to find better TEMPO analogs. TEMPO is a poor laccase substrate, mainly due to its slow kcat (∼10 s−1) [183]. Oxidized TEMPO can be degraded/deoxygenated to the inactive 2,2,6,6-tetramethylpiperidine, significantly reducing the catalytic turnover efficiency. The problem may be addressed by systematically evaluating piperidine/pyrrolidine nitroxyls and other TEMPO analogs [144, 144a, 181]. A dual-mediator system may be used, in which a good laccase substrate could quickly mediate the oxidation/recycling of TEMPO by the laccase, while the TEMPO oxoammonium effectively oxidizes the target [125, 183], although likely at higher cost. In addition to TEMPO and analogs, polyunsaturated hydrocarbons (including fatty acids/lipids) and derivatives may also be applied in laccase catalysis. In linoleate, the “doubly” allylic C site, flanked by two >C¨ C< bonds, may form an allylic C• readily after a C—H bond homolysis (by H-abstraction). The radical may be oxygenated to a peroxy radical or delocated to another C site (via >C¨ C< bond rearrangement), and these radicals or their descendants may serve as potent oxidants. A mediated laccase oxidation may be used to generate the allylic radicals, which may in turn oxidize a target substance. An example of such a dual-mediator laccase biocatalysis has been reported, in which HBT and linoleate mediate a laccase-catalyzed oxidative degradation of phenanthrene [129]. The effect might also contribute to the often observed benefit (for laccase biooxidation) from Tween 80 or other surfactants containing unsaturated moieties. It has long been suspected that some of the natural delignification microbial systems employ endogenous mediators for their laccases. For Pycnoporus cinnabarinus, the native metabolite 3-hydroxyanthranilic acid has been shown to act as a laccase mediator (see Section 2.2.1.7) [133]. For Petriellidium fusoideum, a 1.6 kDa Fe-containing compound has been speculated to assist its extracellular laccase

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[186]. Identifying naturally occurring mediators is of interest for the development of highly efficient, inexpensive, and safe laccase biocatalysis systems [132]. 2.3.1.3 Cooperation with Other Enzymes Targets of laccase catalysis are often embedded in, entangled with, or shielded by heterogeneous substances inert to laccase action. Other enzymes may be required to transform or degrade those substances to expose the targets for laccase. For instance, a mixture of cellulase, hemicellulase, and laccase may be used to synergistically degrade lignocellulosic biomass. Laccase may also be used to support the catalysis of other enzymes for in situ cosubstrate generation, cofactor recycling, or inhibitor elimination. For instance, a laccase-catalyzed phenol → (semiquinone →) quinone oxidation could recycle the reduced quinone which is used to generate active Fenton reagents (oxy radicals), in collaboration with aryl alcohol oxidase or other enzymes [7, 187]. Laccase could also be applied to oxidize Mn2+ to Mn3+ , whose interaction with O2 could then (via O2− ) generate H2O2 for feeding Mn-peroxidase [177]. A laccase subsystem may even serve as the fi nal electron acceptor for flavoenzymes (see Section 2.3.2.4). 2.3.2 New Leads for Laccase Application

Besides the laboratory and commercialized applications discussed in Sections 2.2.1 and 2.2.2, other potential laccase catalyses have been recently proposed or reinvestigated, as the field continues to expand and progress. 2.3.2.1 Laccase-Based Defense Against Biological and Chemical Warfare Agents The 2001 anthrax letter cases in the USA, and the 1995 sarin attack in Tokyo illustrate the threat that biological and chemical weapons pose when used by terrorists. They also emphasize how important it is for security, emergency, and medical agencies to be adequately equipped to decontaminate large, open areas such as subways, airports, and other public facilities when attacked by biological or chemical warfare agents. Traditionally used decontamination systems are not suited for such tasks, as they generally involve chlorine-based chemistry, and their corrosiveness can cause bodily harm as well as material damage, particularly on sensitive or electronic equipments. Development of new, milder, and more universally applicable decontamination systems is highly desirable. Industrial oxidoreductases (such as laccases) hold promise because of their ability to oxidize a wide range of harmful substances, including biological warfare agents and nerve agents like VX and anthrax spores. A laccase system could offer considerable advantages over classical types of decontaminants. Laccases are capable of catalytically transforming substrates, in amounts many times the enzymes’ own weight, in seconds or minutes. Since most known laccases function optimally at pH values near neutrality, there should be few compatibility or corrosion concerns. Also flammability concerns would be minimal. In addition, a laccase-based decontaminant should pose little health or environmental danger and leave no or


little hazardous byproducts. A laccase-based decontaminant might be benign enough to be used directly on the exposed/injured skin. Another major advantage is that a laccase-based decontaminant could be provided as dry pellets or granulates suitable to a carrier-system designed for decontamination. Nerve agents such as organophosphorus compounds are strong inhibitors of acetylcholinesterase, which binds and hydrolyzes the neurotransmitter acetylcholine. The inhibition can result in convulsions, salivary secretion, behavioral incapacitation, muscle weakness, and ultimately death due to respiratory failure. The application of hydrolytic enzymes to detoxify organophosphorus-based chemical nerve agents has been under investigation for decades, as the huge stockpiles of about 200 000 tonnes of nerve agents worldwide is a great international concern. Since the early work on the ability of mammalian tissue to hydrolyze diisopropyl fluorophosphates, a variety of hydrolytic enzymes with activity on organophosphorus compounds have been identified, purified, and characterized from sources such as mammals, cephalopods, and microorganisms. Reports on oxidative degradation of nerve agents by enzymes are more recent. A fungal laccase has been demonstrated to be capable of degrading VX [188], similar to the action of chloroperoxidase [188a]. To effectively degrade phosphorothioate-based chemical nerve agents (cf. Table 2.4) is not trivial. Many phosphorothioates are either resistant towards hydrolysis, or the hydrolysis products are very toxic. Laccasemediated oxidation, in contrast, may convert these phosphorothioates into less toxic products, providing an alternative way to degrade chemical warfare agents [188]. Because phosphorothioates are also present in many insecticides (such as parathion or paraoxon), laccase-mediated oxidation may be applied as a tool to remediate polluted water reservoirs or ponds. To further develop this application, the current over-stoichiometric levels mediator dosing needs to be reduced by finding more effective mediators or conditions. In contrast to the amount of published data on using enzymes to degrade various chemical warfare agents, few reports exist on applying enzymatic decontamination or inactivation of biological warfare agents, such as anthrax spores. Increasing research and development activities are ongoing, however. For instance, Novozymes recently collaborated with the Defense Advanced Research Projects Agency of USA to demonstrate the efficiency of vanadium chloroperoxidase and laccase in killing Bacillus anthracis spores. The preliminary results suggest that a mediated laccase system could play a significant part in a future benign decontaminant and close the current decontamination gap against biological agents such as B. anthracis spores. 2.3.2.2 Degradation of PAH, Plastics, or Lipids With the advent of mediated laccase catalysis, many molecules that are not direct laccase substrates may be oxidatively transformed by laccase-driven reactions. Among the potential laccase oxidation targets, PAHs are attracting increasing attention, due to their environmental presence as fossil fuel pollutants or other industrial wastes, as well as their recalcitrance to conventional clean-up techniques. Laccases have recently been used to transform various PAHs [189–191].

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In general, the Eº of these compounds is too high for laccases to directly oxidize them. The use of mediators may allow indirect electron transfer, H-abstraction, or other reactions to take place, making laccase competitive with other enzyme catalysts such as cytochrome P450 or peroxidases. Reported laccase-targetable PAHs include naphthalene, anthracene, phenanthrene, pyrene, benzo[a]pyrene, fluorene, fluoranthene, dibenzothiophene, and carbazole [129, 132, 135, 191–193]. Often a PAH is oxidized to quinone, and an S-containing PAH to sulfoxide. PAHs may also be induced to polymerize either among themselves or with non-toxic substances (such as humic materials), thus facilitating easy removal by means such as adsorption, sedimentation, or filtration. Among the mediators studied, N—OH types are generally more effective than ABTS or phenothiazine type. Often the extent of PAH oxidation does not depend on the PAH’s ionization potential [191, 192, 194], consistent with the H-abstraction mechanism. In most reported studies, higher than stoichiometric amounts of mediators are needed, indicating that the process could be further improved. Using organic cosolvents in combination with immobilized laccase may be beneficial to enhance the solubility of highly hydrophobic PAHs [195]. Laccases may also be expressed and secreted from the roots of transgenic plants, whose growth in contaminated soil could allow in situ laccase biooxidation/phytoremediation [196]. Mediated laccase oxidation processes may be applied to degrade olefinic, polyethylene, polyurethane, or polyamide plastics, and are thus potentially useful for many biodegradation applications [197–199]. It is likely that laccase-oxidized mediators (such as the N—O• form of HBT) initiate radical reactions, leading to reduced polydispersity, shortened chain elongation, weakened tensile strength, or overall disintegration of the plastics. Recently, laccase has been shown capable of oxidizing and degrading lipids such as trilinolein and methyl linoleate [200, 201]. These unsaturated fatty compounds are not typical laccase substrates, but their allylic C—H groups are susceptible to laccase-driven, mediated H-abstraction (see Section 2.3.1.2). Laccase-oxidized N—O• mediators can abstract the H to form a pentadienyl linoleic radical, which then reacts with O2 to yield subsequently a peroxy radical, a hydroperoxide, or an epoxide. The reaction is of interest because of the occurrence of the fatty compounds in wood and food, which may get involved in laccasecatalyzed delignification and food modification, respectively. 2.3.2.3 Enzymatic Fuel Cells/Batteries Fuel cells are very attractive energy sources, particularly at portable scale, with high energy efficiency, low pollution, and desirable refueling convenience [202]. A fuel cell uses two electrodes to reroute the electron flow in a redox reaction: an anode to take electrons from a reducing substance (like H2, carbohydrate, or alcohol), and a cathode to transfer the electrons, after carrying out work in the circuit, to an oxidizing substance (like O2 ). As a biocatalyst, laccase may be adsorbed, entrapped, or wired onto the cathode to greatly diminish the overvoltage for the four-electron cathodic O2 reduction. How to keep the laccase active and to enhance the heterogeneous/interfacial electron exchange (via mediation, chemical modification, or protein engineering) would be important for the devel-


opment of the application [203, 204]. Cathodic laccase may be joined by an anodic oxidoreductase (like carbohydrate oxidase or alcohol dehydrogenase) and/or H + transporting oxidoreductase for a “complete” enzymatic fuel cell or battery [205, 206]. In addition to fuel cells, laccase-driven bioelectrochemical cells may be applied in artificial photosynthesis [207]. 2.3.2.4 Novel Synthetic Applications Using various mediators, laccase may drive new, “green” synthetic applications in addition to those discussed in Section 2.2.1.5 [2]. Laccase-oxidized N—O• mediators may abstract H from allylic C—H to form allylic C•, which may result in the homolysis of adjacent C—O, C—H, or C—C bonds, or in the rearrangement of C¨ C bonds. Under laccase oxidation, benzylic or aliphatic allylic hydrocarbons may be converted to benzylic or allylic alcohol, alkene, or cyclized products [2, 145, 156, 208, 222]. Toluene, benzyl alcohol, or benzyl amine may be oxidized to benzaldehyde; N-hydroxyamine may be oxidized to oximes, nitrones, nitriles, or carboxylic acids; and propargyl alcohol may be oxidized to the corresponding carbonyl [141, 142, 145, 209–211]. Unsaturated fatty acids or lipids may be converted to hydroperoxide or epoxide derivatives amenable for further functionalization [200, 212]. Similarly, some alkenes (such as 1-(3,4dimethoxyphenyl)-prop-1-ene) may be transformed by laccase-driven epoxidation, hydroxylation, or other reactions [213]. Laccase-oxidized TEMPO (oxoammonium form) may oxidize primary alcohols via an ionic oxidation mechanism, as discussed in Section 2.3.1.2 [107, 145]. This process could be of special interest for carbohydrate targets. For instance, the 6-OH of glucose may be oxidized to an aldehyde or even a carboxyl by TEMPO-mediated laccase catalysis [214, 215], a reaction potentially useful for modification/functionalization of oligo/polysaccharides (like cellulose or amylose), as well as synthesis of bioactive glycosides or other carbohydratebased compounds [107, 216, 217]. For these biocatalytic syntheses, laccase basically serves as a generator of diffusible and active radical or cationic species, which then act on targets chemically to carry out or induce peroxygenation, hydroxylation, epoxidation, C—C cleavage, decarboxylation, C¨ C rearrangement, or other reactions. The mediation may impart a higher degree of regioselectivity to the laccase-catalyzed process (such as acting on primary alcohols instead of secondary alcohols). But the enantioselectivity, often sought after for high-value syntheses, is in general low for mediated laccase biocatalysis, although the TEMPO mediation may have a moderate stereo-preference due to the involvement of the tetrahedral transition-state intermediate (cf. Section 2.3.1.2). Future designer mediators might be able to address this shortcoming [141, 142, 145, 209]. Nevertheless, as laccases are able to use inexpensive O2 as the cosubstrate/electron acceptor, instead of expensive NAD(P) + or other cofactors, and to generate harmless H2O as the coproduct, they should be highly desirable as synthesis catalysts. TEMPO, N-hydroxyphthalimide, and other compounds are well known for their function in organic synthesis [180]. The ability of laccases to regenerate these compounds for catalytic turnover may allow the exploitation of novel mediated laccase biooxidations.

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Mediated laccase oxidation has recently been applied to reoxidize/turnover flavoenzymes, such as cellobiose dehydrogenase and pyranose-2-oxidase [138, 218], during their catalysis. Replacing the flavoenzymes’ native, H2O2-producing interaction with O2, laccase-oxidized mediators may take the electrons from the reduced flavin, shuttle them to laccase, which then transfer them to O2 to produce H2O. Although the reported studies were aimed primarily at reducing the flavoenzyme-destabilizing H2O2 production, the approach may be applied to other oxidoreductases dependent on NAD(P) + , cytochrome c, or other non-O2 cofactors. Some of the enzymes have desirable stereoselective biocatalysis, but are difficult to develop as industrial biocatalysts due to their dependence on expensive cofactors. 2.3.2.5 Biorefinery Biorefinery, or the technology to transform cellulose, hemicellulose, lignin, and other biomass components into value-added materials, is attracting more and more attention for the development of sustainable and environment-friendly energy and other industries. Many cellulases and hemicellulases may be applied to readily hydrolyze the carbohydrates to fermentable “platform” sugars, but the enzymes often encounter unproductive inhibition from lignin [219, 220]. The modification and/or degradation of lignin by laccases, which is suspected to be part of the natural microbial wood degradation, may mitigate the inhibition of the hydrolases. Laccases may also be applied to degrade lignin-derived phenolics that are toxic to ethanogenic yeast for ethanol fermentation [221], or other microbes for other fine chemicals fermentations.

2.4 Further Developing Laccase Catalysis

Currently only two or so bulk, industrial laccase applications have been commercialized (see Section 2.2.2). However, there are reasons to believe that laccases have a bright future as biocatalysts. Compared with conventional chemical catalysts, laccases may require less energy, water, or other resources while discharging less waste. Compared with other potential enzyme catalysts, laccases may require simpler protein/cofactor biosynthesis and rely on less expensive cosubstrates. The scope of laccase-catalyzable reactions may be significantly expanded with the help of redox mediators. To further develop various promising application leads, many optimizations need to be made. For the initial and laboratory-level research, improving the reactivity and stability of laccase and its production are of particular importance. 2.4.1 Laccase Engineering

Limited protein engineering on laccase, by site-directed mutagenesis as well as directed evolution, has been previously made to probe the enzyme’s structure–

2.4 Further Developing Laccase Catalysis

function relationship [23–27, 151, 204]. Since this initial work, our knowledge on the atomic structure (see Section 2.1.1) and natural diversity (see Section 2.3.1.1) of laccase has been significantly expanded, which should enable more effective rational design and gene shuffling-based protein engineering of laccase. For instance, a recent site-directed engineering aimed at an aspartate important for the substrate binding/activation has led to significant shiftings of the optimal pH of a Trametes laccase [231]. With regard to its reactivity, laccase’s T1 Eº is probably the most important factor. It is known that both the high- and low-Eº laccases can have the same amino acid residues to ligate the T1 Cu. Mutating the axial ligand has not been able to convert a low-Eº laccase into a high-Eº laccase [23, 25–27]. A random mutagenesis or gene shuffling approach may be needed to alter the global conformation of the protein backbone to perturb the T1 Cu ligation geometry enough to significantly raise the Eº. As discussed in Section 2.3.1.1, some bacterial or archaeon laccases have very high alkaline pH or thermal stability. Understanding the structural basis of the property may guide us to render fungal laccases, which generally are more active at ambient temperature or neutral pH than bacterial ones, more alkaline or thermal activity/stability desirable for many applications. Further studying the laccases from marine microorganisms or extremophiles may help us to enhance the salinity tolerance or other robustnesses of the biocatalyst [223]. Directed evolution may be used to engineer laccases active in organic solvents [24, 224]. Target-specific/binding molecules may be fused with or linked to laccase to create “smart” biocatalysts. For example, llama heavy-chain antibody may enhance the specificity of laccase towards a target that is immunogenic to the antibody [224a]. Carbohydrate-binding motifs may bring laccase to close vicinity of cellulose-adsorbed targets for better action [224b]. Peroxisome-targeting signal may insert laccase to a fatty particulate for better oxidation [225]. Artificial binding peptides, constructed by either organic synthesis (such as combinatorial chemistry) or genetic engineering (such as phage display), may also enhance laccase’s specificity, such as that towards carotenoid [73]. 2.4.2 Laccase Production

Currently, laccase production in fermentative microbes is the most attractive and feasible method, due to its proven capability and reliability, plus it is constantly being improved by advances in genetic engineering, molecular biology, and bioreactor engineering. Alternative technologies have certain potentials, but they remain largely exploratory at present. Among them transgenic plants are emerging as promising production hosts for laccase [226–229]. Transgenic plant production may be advantageous over microbial production, since the former would diminish potential microbial toxin contamination. Plant-expressed laccase may be used for in situ phyto-biocatalysis [196, 230]. Economically, the production process is easily scalable and could use the existing agriculture infrastructure for

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cultivation, harvest, storage, and processing, resulting in capital investment saving. However, extracting and purifying the expressed enzymes may be costly.

Acknowledgments

We thank Henrik Lund, Joel R. Cherry, Hanne H. Pedersen, Robert L. Starnes, and Ejner B. Jensen of Novozymes for critical reading and suggestions.

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Miyakoshi, T. J Mol Catal, B Enzym 2000, 8, 213–219. 214 Jetten, J.M., van den Dool, R., van Hartingsveldt, W., Besemer, A.C. US patent US6716976, 2004. 215 Viikari, L., Kruus, K., Buchert, J. International patent WO 99/23117, 1999. 216 Monti, D., Candido, A., Silva, M.M.C., Ken, V., Riva, S., Danieli, B. Adv Synth Catal 2005, 347, 1168–1174. 217 Baratto, L., Candido, A., Marzorati, M., Sagui, F., Riva, S., Danieli, B. J Mol Catal, B Enzym 2006, 39, 3–8. 218 Baminger, U., Ludwig, R., Galhaup, C., Leitner, C., Kulbe, K.D., Haltrich, D. J Mol Catal, B Enzym 2001, 11, 541– 550. 219 Berlin, A., Balakshin, M., Gilkes, N., Kadla, J., Maximenko, V., Kubo, S., Saddler, J. J Biotechnol 2006, 25, 198–209. 220 Xu, F. In: Ohmiya, K., Sakka, K., Karita, S., Kimura, T., Sakka, M., Onishi, Y., eds. Biotechnology of Lignocellulose Degradation and Biomass Utilization. Tokyo: Uni Publishers, 2004, pp. 793–804. 221 Larsson, S., Cassland, P., Jönsson, L.J. Appl Environ Microbiol 2001, 67, 1163–1170. 222 Nicotra, S., Intra, A., Ottolina, G., Riva, S., Danieli, B. Tetrahedron Asymmetry 2004, 15, 2927–2931. 223 d’Souza, D.T., Tiwari, R., Sah, A.K., Raghukumar, C. Enzyme Microb Technol 2006, 38, 504–511. 224 Alcalde, M., Bulter, T., Zumarraga, M., Garcia-Arellano, H., Mencia, M., Plou, F.J., Ballesteros, A. J Biomol Screen 2005, 10, 624–631. 224a van der Logt, C.P., Parry, N.J., Howell, S., Little, J. Method of delivering a benefit agent. US patent US6642196B2, 2003. 224b van der Logt, C.P., Howell, S., Little, J., Hemmington, S., Parry, N.J., Smith, R. G. Method of treating fabrics and apparatus used therein. US patent US6586384B2, 2003. 225 Howell, S., Little, J., van der Logt, C.P.E., Parry, N.J. International patent WO200146364-A2, 2001.

References 226 Howard, J.A., Hood, E., Jilka, J. US Howard, J.A., Hood, E.E. In Vitro Cell patent US6800792B1, 2004. Dev Biol 2002, 38, 62A. 227 Bailey, M.R., Woodard, S.L., Callaway, 229 Chen, X., Wang, G. Chinese patent E., Beifuss, K., Delaney, D., CN1607249-A, 2005. Magallanes-Lundback, M., Lane, J., 230 Sonoki, T., Kajita, S., Ikeda, S., Uesugi, Horn, M.E., Ward, M., van Gastel, F., M., Tatsumi, K., Katayama, Y., Iimura, Howard, J.A., Hood, E.E. Appl Y. Appl Microb Biotechnol 2005, 67, Microbiol Biotechnol 2004, 63, 390–397. 138–142. 228 Ruby, K.L., Bailey, M.R., Woodward, 231 Madzak, C., Mimmi, M.C., Caminade, S.L., Callaway, E., Delaney, D., Beifuss, E., Brault, A., Baumberger, S., Briozzo, K., Magallanes-Lundback, M., Lane, J., P., Mougin, C., Jolivalt, C. Protein Eng Horn, M.E., Ward, M., van Gastel, F., Des Sel 2006, 19, 77–84.

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3 Biocatalytic Scope of Baeyer–Villiger Monooxygenases Marco W. Fraaije and Dick B. Janssen

3.1 Introduction 3.1.1 The Baeyer–Villiger Reaction

A commonly used oxidation reaction in organic synthesis is the conversion of carbonylic compounds into the corresponding esters or lactones. This reaction was discovered in 1899 by Adolf Baeyer and Victor Villiger and therefore is called the Baeyer–Villiger reaction [1]. The Baeyer–Villiger reaction is typically performed using a peroxyacid, resulting in the insertion of an oxygen atom between the carbonyl carbon and a neighboring carbon atom. For several decades the exact mechanism by which this oxygenation reaction proceeds was debated [2]. The reaction mechanism was finally elucidated in 1953 when Doering and Dorfman studied the Baeyer–Villiger oxidation of 18O-labeled benzophenone [3]. Upon oxidation of this ketone with perbenzoic acid, the labeled oxygen atom ended up as the carbonyl oxygen of the ester that was formed. This outcome confirmed a mechanism that was first proposed by Criegee [4] and involves formation and decay of a so-called “Criegee intermediate.” This tetrahedral intermediate is formed by a nucleophilic attack of the peroxyacid onto the ketone (Fig. 3.1). Subsequent decay of this labile intermediate involves migration of a substituent from the carbonyl carbon, yielding an ester or lactone. The Baeyer–Villiger reaction is widely appreciated in organic synthesis as it is applicable to a broad range of carbonylic compounds while the moiety that will migrate during the reaction can also be predicted to some extent. Unsubstituted groups (e.g. a methyl) adjacent to the carbonyl function represent poor migrating groups while highly substituted groups (e.g. tertiary alkyl) have a good migratory aptitude. An additional attractive feature of the Baeyer–Villiger reaction is that the migrating group will typically retain its configuration. This allows effective and selective oxidation reactions. However, a major drawback of the reaction is the intrinsic need for a potent and therefore hazardous oxidizing agent. The classical approach to perform a Baeyer–Villiger reaction includes the use of organic

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3 Biocatalytic Scope of Baeyer–Villiger Monooxygenases

Fig. 3.1 The Baeyer–Villiger reaction proceeds via formation and subsequent rearrangement of a Criegee intermediate.

peroxyacids as catalyst. In addition to the fact that these reactive compounds have to be handled with care, they are also relatively expensive. This has triggered development of more gentle catalytic systems that enable Baeyer–Villiger reactions, such as the use of hydrogen peroxide-dependent catalysts (see [5] for a recent review). An obvious alternative “green chemistry” approach would be the use of a biocatalyst that is able to catalyze Baeyer–Villiger reactions. 3.1.2 Baeyer–Villiger Biocatalysts: Classification and Occurrence

The first indications for the natural existence of so-called Baeyer–Villiger monooxygenases (BVMOs) were reported in the late 1940s. It was observed that certain fungi were able to convert steroids via a Baeyer–Villiger reaction [6]. In 1965 the first BVMO was purified from a Pseudomonas sp. [7]. In the last few decades a number of other BVMOs have been purified and characterized. This has revealed the existence of two discrete classes of BVMOs [8]: types I and II. 3.1.2.1 Type I Baeyer–Villiger Monooxygenases Type I BVMOs are composed of only one polypeptide of about 500 amino acids. They contain FAD as flavin cofactor and require NADPH and molecular oxygen for activity [9]. These monooxygenase are typically soluble proteins located in the cytosol of bacteria or fungi. Most of the presently known BVMOs are of this type. Type I BVMOs are sequence related to two other flavoprotein monooxygenase classes: hetero-atom flavin-containing monooxygenases and N-hydroxylating flavin-containing monooxygenases [10]. This reflects a shared reactivity as all three flavoprotein classes employ a FAD cofactor to perform oxygenation reactions. 3.1.2.2 Type II Baeyer–Villiger Monooxygenases Type II BVMOs [8] are typically composed of two different subunits in which a reductase subunit utilizes NADH to reduce FMN while the other subunit utilizes this reduced flavin to perform a Baeyer–Villiger reaction. While type I BVMO genes have been frequently reported, cloning of a type II BVMO gene has not been described in the literature. For a long time only several sequenced N-termini were known [8] which prohibited classification of these monooxygenases in the flavoprotein suprafamily. Recently, however, a sequence of a “limone monooxy-

3.1 Introduction

genase” (GI12054950) has been deposited in the sequence database which shares homology with the known type II BVMO N-termini. This bacterial monooxygenase gene, which is part of a gene cluster involved in limonene degradation, probably represents the oxygenase component of a type II BVMO that is involved in the conversion of 1-hydroxy-2-oxolimonene or a related terpene [22]. Based on the inspection of the published N-terminal sequences and database searches using the fully sequenced putative type II BVMO, it can be concluded that type II BVMOs are rather rare enzymes. Only 10 sequence homologs can be identified, which is in sharp contrast with more than 400 type I BVMO genes that can be found by in silico inspection of sequenced genomes. In addition to the corresponding oxygenase subunit genes, a reductase component is also necessary to obtain a functional type II BVMO system. This clearly complicates the cloning and expression of type II BVMOs and might partly explain the lack of type II BVMOs in a recombinant form. 3.1.2.3 Alternative Baeyer–Villiger Biocatalysts Recently several enzymes have been discovered that display Baeyer–Villiger activity although they do not resemble the above mentioned BVMOs. A monooxygenase (MtmOIV) involved in the biosynthesis of mithramycin, an antitumor drug, was shown to catalyze a Baeyer–Villiger reaction [23]. Sequence analysis has shown that this flavoprotein cannot be classified as a type I or II BVMO as it is sequence related to flavin-containing monooxygenases belonging to another flavoprotein family (for a review on the classification of flavoprotein monooxygenases, see [10]). This suggests that at least one other class of flavin-dependent BVMOs exists (type III BVMOs) of which MtmOIV is the first identified member. Other reports have indicated that some specific cytochrome P450s are capable of modifying steroids via a Baeyer–Villiger reaction [24, 25]. These heme-containing monooxygenases are supposed to exploit the hydroperoxy heme intermediate (FeIII-OOH) as a “peracid catalyst.” Future studies will reveal more mechanistic details concerning these newly identified Baeyer–Villiger biocatalysts. Some hydrolases can also be used to catalyze Baeyer–Villiger oxidations [26]. It has been shown that, analogous to cofactor-free haloperoxidases, serine hydrolases are capable of forming peroxyacids from carboxylic acids upon consumption of hydrogen peroxide. However, such a catalytic system lacks enantio-, regio-, and/or chemoselectivity as the hydrolase merely acts as a peroxyacid-forming catalyst. A hydrolase does not offer a suitable active site for the peroxyacid and/or substrate to bind. As a consequence the reaction of the formed peroxyacid with the carbonylic substrates will take place in solution without the steric confinement of an enzyme active site, prohibiting effective selectivity. Only in one specific case it has been reported that a lactone could be obtained with a moderate enantiomeric excess (21%) using lipase B from Candida antarctica [27]. This selectivity can be explained by the fact that the chiral carboxylic acid itself also contained a carbonylic function allowing an intramolecular Baeyer–Villiger reaction upon formation of the peroxyacid. The observed enantioselectivity is presumably caused

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by the preferred oxidation of one of the ketoacid enantiomers. Except for a lack of selectivity, another disadvantage of the use of (per)hydrolases to perform Baeyer–Villiger reactions is their ability to hydrolyze the formed esters or lactones.

3.2 Type I Baeyer–Villiger Monooxygenases: Versatile Oxidative Biocatalysts

While alternative Baeyer–Villiger biocatalysts have been reported, type I BVMOs appear to be the most convenient enzymes to perform Baeyer–Villiger reactions. This is reflected in the large number of reports in which applications based on these monooxygenases are demonstrated (for a recent review see [28]). In the next paragraphs, the biocatalytic scope of type I BVMOs will be reviewed. First, the mechanism by which these enzymes are able to perform Baeyer–Villiger reactions is discussed. The mechanistic data and enzyme engineering efforts will be discussed in relation to the recently elucidated structure of a BVMO. Furthermore, the catalytic potential will be highlighted by elaborating on the kinetic properties and substrate specificities of BVMOs. Also recent advances in knowledge and technical approaches concerning coenzyme regeneration and biocatalyst stability are touched upon. 3.2.1 Mechanistic and Structural Properties of Type I BVMOs

The best-studied BVMO is cyclohexanone monooxygenase (CHMO) from Acinetobacter sp. NCIMB 9871. This type I BVMO was first isolated in 1976 [29] which set off many biocatalytic studies involving this monooxygenase [28]. Over the years a large number of different substrates (>100) have been reported for this bacterial monooxygenase. The enzyme has been used as prototype BVMO to show that these enzymes are not only capable of performing Baeyer–Villiger oxidations but can also oxygenate sulfides, selenides, amines, phosphines, olefins, and iodideand boron-containing compounds. In a seminal paper in 1982 the first pre-steadystate kinetic analysis of CHMO was described [30]. Using the stopped-flow technique it was shown that CHMO is able to form and stabilize an oxygenated form of the bound FAD cofactor: a C4a-peroxy-FAD (Fig. 3.2). This intermediate is formed after reduction of the flavin cofactor by NADPH, which in turn triggers a fast reaction with molecular oxygen yielding C4a-peroxy-FAD. It was shown that external addition of hydrogen peroxide to CHMO does not result in formation of this peroxyfl avin intermediate [30]. Apparently the enzyme is unable to use hydrogen peroxide and can only form the reactive intermediate by using the NADPH coenzyme and molecular oxygen. Substrate binding has no effect on the formation of the oxygenated flavin cofactor. Such a seemingly uncontrolled consumption of valuable NADPH and formation of a reactive enzyme species is not observed other monooxygenase systems.


Fig. 3.2 Catalytic mechanism of type I BVMOs.

In cytochrome P450 monooxygenases and FAD-containing hydroxylases, formation of the oxygenating enzyme intermediate is tightly coupled to substrate binding, which prevents unproductive coenzyme consumption. In CHMO, such an uncontrolled consumption of NADPH (uncoupling) is largely prevented by the effective stabilization of the peroxyflavin intermediate. In the presence of NADPH and the absence of a suitable substrate only a very slow rate of hydrogen peroxide formation is observed for CHMO. For phenylacetone monooxygenase (PAMO) and 4-hydroxyacetophenone monooxygenase (HAPMO) very slow uncoupling reactions have also been observed, with rates lower than 0.1 s−1 (M.W. Fraaije unpublished results). The reactive peroxyflavin enzyme intermediate mimics the function of a peroxyacid as it can perform a nucleophilic attack on the carbonyl carbon of a ketone or aldehyde resulting in a Baeyer–Villiger oxidation. In analogy with peroxyacidcatalyzed Baeyer–Villiger reactions, the reaction would involve formation of a flavin-substrate Criegee intermediate in the active site of CHMO. Decay of this intermediate results in formation of the ester product. While the oxygenated C4a-peroxy-FAD intermediate has been observed in CHMO [30, 31], no Criegee-intermediate has ever been observed in pre-steady-

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state kinetic studies, indicating that this is not a long-lived enzyme species. In case of CHMO it has been found that release of the ester product precedes decay of the hydroxyflavin that is formed upon collapse of the Criegee intermediate [30] (Fig. 3.2). The catalytic cycle is completed by release of NADP+ . A recent kinetic study of CHMO has revealed that release of the oxidized coenzyme, NADP+ , is limiting the rate of catalysis [31]. This indicates that the conversion rate is not always determined by the nature of the substrate or product and is in line with the observation that other substrates display similar steady-state kinetic parameters when compared with cyclohexanone. A similar kinetic behavior has been observed for eukaryotic flavin-containing monooxygenases which are distantly related in sequence [32]. In our laboratory we have observed that HAPMO and PAMO share several kinetic properties with CHMO (unpublished results). With these BVMOs also the peroxyflavin is formed and stabilized in the absence of a substrate, no Criegee intermediate is observed and NADP+ has been found to represent a competitive inhibitor with respect to NADPH. Recently we have been able to demonstrate by using electrospray ionization mass spectrometry (ESI-MS) that the coenzyme NADPH/NADP+ is bound to the enzyme throughout the catalytic cycle [33]. In 2004 the first crystal structure of a BVMO, PAMO, was published [34]. This BVMO represents a thermostable enzyme as it originates from the mesothermophilic bacterium Thermobifida fusca [35]. Substrate profi ling studies have shown that the enzyme prefers aromatic substrates while it also accepts some aliphatic substrates. The crystal structure of PAMO was solved at 1.7 Å resolution and revealed the presence of two dinucleotide binding domains and a helical subdomain (Fig. 3.3). The dinucleotide binding domain formed by residues from the N- and C-termini binds the FAD cofactor in an extended conformation. Such a mode of FAD binding has been observed for many other flavoproteins. The combination of an FAD-binding domain with an NADPH-binding domain is also reminiscent of other FAD-containing oxidoreductases. Contrarily, the helical domain that flanks both cofactor-binding domains and which forms part of the active site appears to be unique to type I BVMOs. Unfortunately, we have been unable to obtain crystals that contain a bound NADPH/NADP+ or a substrate/product which would reveal the binding mode of these ligands. Nevertheless, binding of NADPH could be easily modeled into the structure using the observed binding modes in other protein structures as a guide. This modeling exercise has suggested that two conserved basic residues, R217 and K336, are crucial for recognition of the adenine and 2′-phosphate moieties of NADPH, respectively. Elucidation of the structure of PAMO enabled identification of an active site residue that appears to be key to catalysis in type I BVMOs: R337 (Fig. 3.3). R337 is located at the re side of the flavin and points towards the reactive part of the FAD cofactor. This residue is strictly conserved in all type I BVMOs. Replacing R337 in PAMO (unpublished results) and HAPMO by an alanine yielded inactive


Fig. 3.3 Crystal structure of phenylacetone monooxygenase from Thermobifida fusca. The FAD cofactor and the active site arginine R337 are shown in sticks. The Cα atoms of residues that are discussed in the text are shown in spheres.

enzymes [37]. Based on the structure of PAMO we have proposed that the positive charge of this arginine may be crucial (a) to stabilize the negatively charged peroxy-FAD intermediate and/or (b) to promote and stabilize the negatively charged Criegee intermediate (Fig. 3.2). The delicate function of this active site residue is also apparent from the absolute strict conservation of this residue within all identified type I BVMO sequences. It is also striking to note that this active site arginine is the only active site residue that is conserved among type I BVMOs. The extensive variation of active site residues might reflect the plasticity in substrate specificities observed for these monooxygenases. As no structure is available of PAMO with a substrate or product bound in the active site, it is difficult to assign residues that form the substrate-binding pocket. In a recent enzyme redesign study it was demonstrated that by deleting two residues in a loop region that is close to R337 the substrate specificity of PAMO can be significantly altered [37]. While PAMO does not accept 2-phenylcyclohexanone, the deletion mutant ΔA441/L442 was found to convert this relatively bulky substrate. It is envisaged that by shortening the loop interacting with R337, the substrate-binding pocket is enlarged and can accommodate bulky substrates (Fig. 3.3). A role of the targeted loop region in substrate binding was substantiated by results obtained from a directed evolution study with CHMO. Screening a library of CHMO mutants created by error-prone PCR resulted in the discovery of three

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single mutants that displayed significantly altered enantioselective behavior [38, 39]. Locating the analogous residues in the PAMO structure reveals that one of them (L443) is part of the abovementioned loop region while the others, Q152 and T393, are in close proximity (Fig. 3.3). These findings confirm that substrates bind at the re side of the flavin in PAMO. A similar situation will hold true for other type I BVMOs as the si side of the flavin in PAMO is occupied by the protein and the corresponding residues are highly conserved among type I BVMOs. 3.2.2 Diversity

Recently, a protein sequence motif has been identified which facilitates identification of putative BVMO genes [11]. This enables effective screening of the available (genome) sequence databases. We have surveyed the presently available sequenced microbia genomes (607) which yielded just over 400 putative type I BVMO sequences. This number is in sharp contrast with the limited number of BVMOs available in recombinant form. It is also striking to note that all identified genes originate from bacteria and fungi. Figure 3.4 shows all known type I BVMO sequences together with the type I BVMO sequences obtained from fully sequenced microbial genomes and the metagenome sequence database in a dendrogram representation. This illustrates the clustering of several sets of sequences in a restricted number of clades. Clustering of several known BVMOs with similar substrate specificities in specific clades suggests that these groups represent BVMOs that act on similar substrates. For example, cyclododecanone monooxygenase (CDDMO) [12] and cyclopentadecanone monooxygenase (CPDMO) [13] have overlapping substrate specificities and are clustered in a distinct clade of 16 BVMO sequences. All these BVMOs probably represent monooxygenases acting on bulky cyclic aliphatic ketones. As can be seen in Fig. 3.4, a similar situation is seen for cyclopentanone monooxygenases (CPMOs) [14] and most cyclohexanone monooxygenases [15– 17], which form two separate clades. This is in agreement with the observation that CPMOs and CHMOs often display different regio- and enantioselectivities [18, 19]. The clustering of enzymes with similar substrate specificities suggests that sequence analysis can be used as a predictive tool to obtain hints concerning substrate specificities of uncharacterized BVMOs. However, most clades contain only one or even no BVMO sequence of which the biocatalytic properties are known. The clade in which HAPMO [20] is located is quite dispersed and totally new substrate specificities can be expected from sequences that are part of this group of BVMO sequences. There is also a distinct cluster of putative BVMO sequences that entails ethionamide monooxygenase from Mycobacterium tuberculosis [21]. It has been shown that this BVMO is able to catalyze a range of Baeyer–Villiger reactions accepting both aromatic and aliphatic substrates. However, the physiological substrate of this enzyme is as yet unknown.


Fig. 3.4 An unrooted phylogenetic tree of type I BVMO protein sequences. Sequences were retrieved from the NCBI and PEDANT sequence databases. In addition to known BVMOs and putative BVMOs identified in annotated microbial genomes, putative homologs retrieved from the metagenome sequence database of the Sargasso sea (NCBI) are also indicated (dotted lines). All characterized BVMOs and their physiological substrates are indicated: CDDMO, cyclododecanone monooxygenase from Rhodococcus rhodochrous; CPDMO, cyclopentadecanone monooxygenase from Pseudomonas HI-70; CPMO1, cyclopentanone monooxygenase from Comamonas testosteroni; CPMO2, cyclopentanone monooxygenases from Brevibacterium sp. HCU; CHMO1, cyclohexanone monooxygenase from Acinetobacter NCIMB

9871; CHMO2, cyclohexanone monooxygenase from Arthrobacter sp. BP2; CHMO3 and CHMO4, cyclohexanone monooxygenases from Rhodococcus sp. Phi1 and Phi2, respectively; CHMO5, cyclohexanone monooxygenase from Xanthobacter autotrophicus Py2; CHMO6, cyclohexanone monooxygenases from Brevibacterium sp. HCU; STMO, steroid monooxygenase from Rhodococcus rhodochrous; PAMO, phenylacetone monooxygenase from Thermobifida fusca; HAPMO, 4hydroxyacetophenone monooxygenase from Pseudomonas fluorescens ACB; EtaA, ethionamide-activating monooxygenase from Mycobacterium tuberculosis H37Rv. Several BVMOs that are discussed in the text as “unexplored BVMOs” and Mtub5 are also indicated.

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3.2.3 Molecular Features

Only a limited number of type I BVMOs have been purified and characterized. However, as type I BVMOs are related in sequence, all representatives will have similar molecular features. From an inventory of biochemical studies on type I BVMOs [12–15, 20, 21, 35, 40] the following common properties have emerged. Type I BVMOs all contain a tightly bound FAD as single prosthetic group and utilize NADPH as electron donor. They typically exhibit a broad pH optimum for activity with optimal activity at slightly basic pH. Most representatives have a molecular mass of 55–65 kDa per subunit and are monomeric. HAPMO displays slightly different characteristics as it is a dimeric enzyme with subunits of 73 kDa. The larger size is due to an additional N-terminal domain of 140 residues which appears to be important for the stability of this dimeric enzyme [41]. This Nterminal extension is found in 10 out of about 400 type I BVMO sequences that can be identified in the genome databases. Most characterized BVMOs represent soluble cytosolic proteins which are easily expressed in E. coli. Only in the case of ethionamide monooxygenase from Mycobacterium tuberculosis were detergents shown to be necessary to isolate the enzyme [21]. This suggests that this prodrug-activating BVMO is membrane-associated which is in line with the proposed physiological role: metabolism of mycolic acids. This atypical feature did not prevent high levels of expression of this monooxygenase in E. coli. For CHMO it has been shown that coexpression of proteins that assist in protein folding increases the expression level of soluble and active protein [42]. The positive effect of coexpression of DsbA, which is important to create internal disulfide bonds, on expression of active CHMO suggests that this BVMO contains one or more crucial disulfide bonds. This is in line with a structural model of CHMO that we have recently constructed using the PAMO structure. This model suggests a disulfide bond between C516 and C520. This structural feature of CHMO might also explain differences in behavior between CHMO produced in E. coli and CHMO isolated from Acinetobacter sp. NCIMB 9871 [43]. 3.2.4 Kinetic Characteristics

With their respective natural substrates, type I BVMOs display kcat values of 5– 20 s−1 while the apparent affinity is in the micromolar range. With these kinetic features, type I BVMOs display catalytic efficiencies that are typical for oxygenases [44]. Only for ethionamide monooxygenase have rather low kcat values been reported, which can be explained by the fact that the natural substrate for this monooxygenase is as yet unknown. The apparent affinity for molecular oxygen has been shown to be excellent for CHMO [30] and HAPMO (unpublished results) with apparent K M values of 95% enantiomeric excess and with a modest yield of 19%. In a recent follow-up study it was demonstrated, using a series of cyclic aliphatic ketones, that this BVMO exhibits exquisite enantioselective behavior, enabling preparative kinetic resolution of cyclic ketones and production of enantio pure lactones [73]. These resolutions cannot be achieved using one of the other available BVMOs. Therefore, this novel BVMO represents a useful biocatalyst as it extends the biocatalytic repertoire in the BVMO toolbox.

3.3 Concluding Remarks

The number of BVMOs that are available in recombinant form has significantly increased over the last few years. Several close homologs of CHMO and cyclopentanone monooxygenase have been described and exploited for biocatalytic studies revealing new complementing enantio- and regioselective properties. Furthermore, BVMOs have been found that cover totally new substrates ranges (e.g. HAPMO and PAMO acting on aromatic ketones and sulfides [55, 60]) or that display novel selectivities (e.g. a novel BVMO from M. tuberculosis enabling kinetic resolution of bicyclic ketones [72, 73]). These research activities have broadened the biocatalytic scope of BVMO applications. The availability of a large collection of sequence microbial genomes will facilitate cloning and exploration of new BVMOs, thereby expanding the biocatalytic scope of BVMOs even further. Except for the discovery of novel BVMOs and their corresponding catalytic abilities, new approaches have also been developed to perform BVMO-mediated conversions. Using E. coli cells expressing CHMO an asymmetric Baeyer–Villiger biooxidation reaction has been performed at kilogram scale [46]. This convincingly shows that by using whole cells expressing a BVMO of interest selective oxidation reactions on a preparative scale are feasible. Another valuable recent development is the elucidation of a BVMO structure at atomic resolution [34]. The structure has provided valuable insight into the complex mechanism of catalysis mediated by FAD-containing BVMOs. It also allows dedicated enzyme redesign studies which will lead to engineered BVMOs with tailored biocatalytic properties. In view of the abovementioned developments, we anticipate a bright future for BVMO-based biocatalytic applications.

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and characterization of a gene cluster for cyclododecanone oxidation in Rhodococcus ruber SC1. J Bacteriol 2001, 183, 6478–6486. Iwaki, H., Wang, S., Grosse, S., Bergeron, H., Nagahashi, A., Lertvorachon, J., Yang, J., Konishi, Y., Hasegawa, Y., Lau, P.C. Pseudomonad cyclopentadecanone monooxygenase displaying an uncommon spectrum of Baeyer–Villiger oxidations of cyclic ketones. Appl Environ Microbiol 2006, 72, 2707–2720. Iwaki, H., Hasegawa, Y., Wang, S., Kayser, M.M., Lau, P.C. Cloning and characterization of a gene cluster involved in cyclopentanol metabolism in Comamonas sp. strain NCIMB 9872 and biotransformations effected by Escherichia coli-expressed cyclopentanone 1,2monooxygenase. Appl Environ Microbiol 2002, 68, 5671–5684. Chen, Y.C., Peoples, O.P., Walsh, C.T. Acinetobacter cyclohexanone monooxygenase: gene cloning and sequence determination. J Bacteriol 1988, 170, 781–789. Brzostowicz, P.C., Gibson, K.L., Thomas, S.M., Blasko, M.S., Rouviere, P.E. Simultaneous identification of two cyclohexanone oxidation genes from an environmental Brevibacterium isolate using mRNA differential display. J Bacteriol 2000, 182, 4241–4248. Brzostowicz, P.C., Walters, D.M., Thomas, S.M., Nagarajan, V., Rouviere, P.E. mRNA differential display in a microbial enrichment culture: simultaneous identification of three cyclohexanone monooxygenases from three species. Appl Environ Microbiol 2003, 69, 334–342. Mihovilovic, M.D., Rudroff, F., Grötzl, B., Kapitan, P., Snajdrova, R., Rydz, J. Mach, R. Family clustering of Baeyer–Villiger monooxygenases based on protein sequence and stereopreference. Angew Chem Int Ed 2005, 44, 3609–3613. Wang, S., Kayser, M.M., Iwaki, H., Lau, P.C.K. Monooxygenase-catalyzed Baeyer–Villiger oxidations: CHMO versus CPMO. J Mol Catal, B Enzym 2003, 22, 211–218.

References 20 Kamerbeek, N.M., Moonen, M.J.H., van der Ven, J.G.M., van Berkel, W.J.H., Fraaije, M.W., Janssen, D.B. 4Hydroxyacetophenone monooxygenase from Pseudomonas fluorescens ACB. A novel flavoprotein catalyzing Baeyer– Villiger oxidation of aromatic compounds. Eur J Biochem 2001, 268, 2547–2557. 21 Fraaije, M.W., Kamerbeek, N.M., Heidekamp, A.J., Fortin, R., Janssen, D.B. The prodrug activator EtaA from Mycobacterium tuberculosis is a Baeyer– Villiger monooxygenase. J Biol Chem 2004, 279, 3354–3360. 22 van der Werf, M.J., Swarts, H.J., de Bont, J.A.M. Rhodococcus erythropolis DCL14 contains a novel degradation pathway for limonene. Appl Environ Microbiol 1999, 65, 2092–2102. 23 Prado, L., Fernandez, E., Weissbach, U., Blanco, G., Quiros, L.M., Brana, A.F., Mendez, C., Rohr, J., Salas, J.A. Oxidative cleavage of premithramycin B is one of the last steps in the biosynthesis of the antitumor drug mithramycin. Chem Biol 1999, 6, 19–30. 24 Fischer, R.T., Trzaskos, J.M., Magolda, R.L., Ko, S.S., Brosz, C.S., Larsen, B. Lanosterol 14 alpha-methyl demethylase. Isolation and characterization of the third metabolically generated oxidative demethylation intermediate. J Biol Chem 1991, 266, 6124–6132. 25 Swinney, D.C., Mak, A.Y. Androgen formation by cytochrome P450 CYP17. Solvent isotope effect and pL studies suggest a role for protons in the regulation of oxene versus peroxide chemistry. Biochemistry 1994, 33, 2185–2190. 26 Carboni-Oerlemans, C., Dominguez de Maria, P., Tuin, B., Bargeman, G., van der Meer, A., van Gemert, R. Hydrolasecatalysed synthesis of peroxycarboxylic acids: Biocatalytic promiscuity for practical applications. J Biotechnol 2006, 126, 140–151. 27 Lemoult, S.C., Richardson, P.F., Roberts, S.M. Lipase-catalysed Baeyer–Villiger reactions. J. Chem Soc, Perkin Trans I 1995, 89–91.

28 Mihovilovic, M.D., Müller, B., Stanetty, P. Monooxygenase-mediated Baeyer–Villiger oxidations. Eur J Org Chem 2002, 3711–3730. 29 Donoghue, N.A., Norris, D.B., Trudgill, P.W. The purification and properties of cyclohexanone oxygenase from Nocardia globerula CL1 and Acinetobacter NCIB 9871. Eur J Biochem 1976, 63, 175– 192. 30 Ryerson, C.C., Ballou, D.P., Walsh, C. Mechanistic studies on cyclohexanone oxygenase. Biochemistry 1982, 21, 2644–2655. 31 Sheng, D., Ballou, D.P., Massey, V. Mechanistic studies of cyclohexanone monooxygenase: chemical properties of intermediates involved in catalysis. Biochemistry 2001, 40, 11156–11167. 32 Poulsen, L.L., Ziegler, D.M. Multisubstrate flavin-containing monooxygenases: applications of mechanism to specificity. Chem Biol Interact 1995, 96, 57–73. 33 van den Heuvel, R.H.H., Tahallah, N., Kamerbeek, N.M., Fraaije, M.W., van Berkel, W.J.H., Janssen, D.B., Heck, A.J. Coenzyme binding during catalysis is beneficial for the stability of 4hydroxyacetophenone monooxygenase. J Biol Chem 2005, 280, 32115–32121. 34 Malito, E., Alfieri, A., Fraaije, M.W., Mattevi, A. Crystal structure of a Baeyer– Villiger monooxygenase. Proc Natl Acad Sci USA 2004, 101, 13157–13162. 35 Fraaije, M.W., Wu, J., Heuts, D.P.H.M., van Hellemond, E.W., Spelberg J.H., Janssen, D.B. Discovery of a thermostable Baeyer–Villiger monooxygenase by genome mining. Appl Microbiol Biotechnol 2005, 66, 393–400. 36 Kamerbeek, N.M., Fraaije, M.W., Janssen, D.B. Identifying determinants of NADPH specificity in Baeyer–Villiger monooxygenases. Eur J Biochem 2004, 271, 2107–2116. 37 Bocola, M., Schulz, F., Leca, F., Vogel, A., Fraaije, M.W., Reetz, M.T. Converting phenylacetone monooxygenase into phenylcyclohexanone monooxygenase by rational design: towards practical Baeyer– Villiger monooxygenases. Adv Synth Catal 2005, 347, 979–986.

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3 Biocatalytic Scope of Baeyer–Villiger Monooxygenases 38 Reetz, M.T., Brunner, B., Schneider, T., Schulz, F., Clouthier, C.M., Kayser, M. M. Directed evolution as a method to create enantioselective cyclohexanone monooxygenases for catalysis in Baeyer– Villiger reactions. Angew Chem Int Ed 2004, 43, 4075–4078. 39 Reetz, M.T., Daligault, F., Brunner, B., Hinrichs, H., Deege, A. Directed evolution of cyclohexanone monooxygenases: enantioselective biocatalysts for the oxidation of prochiral thioethers. Angew Chem Int Ed 2004, 43, 4078–4081. 40 van der Werf, W.F. Purification and characterization of a Baeyer–Villiger mono-oxygenase from Rhodococcus erythropolis DCL14 involved in three different monocyclic monoterpene degradation pathways. Biochem J 2000, 347, 693–701. 41 Kamerbeek, N.M. PhD thesis, University of Groningen, 2004. 42 Lee, D.H., Kim, M.D., Lee, W.H., Kweon, D.H., Seo, J.H. Consortium of fold-catalyzing proteins increases soluble expression of cyclohexanone monooxygenase in recombinant Escherichia coli. Appl Microbiol Biotechnol 2004, 63, 549–552. 43 Secundo, F., Zambianchi, F., Crippa, G., Carrea, G., Tedeschi, G. Comparative study of the properties of wild type and recombinant cyclohexanone monooxygenase, an enzyme of synthetic interest. J Mol Catal, B Enzym 2005, 34, 1–6. 44 van Beilen, J.B., Duetz, W.A., Schmid, A., Witholt, B. Practical issues in the application of oxygenases. Trends Biotechnol 2003, 21, 170–177. 45 Baldwin, C.V., Woodley, J.M. On oxygen limitation in a whole cell biocatalytic Baeyer–Villiger oxidation process. Biotechnol Bioeng 2006, 95, 362–369. 46 Hilker, I., Wohlgemuth, R., Alphand, V. and Furstoss, R. Microbial transformations 59: first kilogram scale asymmetric microbial Baeyer–Villiger oxidation with optimized productivity using a resin-based in situ SFPR strategy. Biotechnol Bioeng 2005, 92, 702–710.

47 Wichmann, R., Vasic-Racki, D. Cofactor regeneration at the lab scale. Adv. Biochem Eng Biotechnol 2005, 92, 225–260. 48 Lundquist, R., Olivera, B.M.: Pyridine nucleotide metabolism in Escherichia coli. I. Exponential growth. J Biol Chem 1971, 246, 1107–1116. 49 de Gonzalo, G., Ottolina, G., Carrea, G., Fraaije, M.W. [Cp*Rh(bpy)(H2O)] 2+ as a coenzyme substitute in enzymatic oxidations catalyzed by Baeyer–Villiger monooxygenases. Chem Commun 2005, 29, 3724–3276. 50 Hollmann, F., Hofstetter, K., Schmid, A. Non-enzymatic regeneration of nicotinamide and flavin cofactors for monooxygenase catalysis. Trends Biotechnol 2006, 24, 163–171. 51 Willetts, A.J., Knowles, C.J., Levitt, M.S., Roberts, S.M., Sandey, H., Shipston, N.F. Biotransformation of endo-bicyclo[2.2.1 ]heptan-2-ols and endo-bicyclo[3.2.0]hept2-en-6-ol into the corresponding lactones. J Chem Soc, Perkin Trans. 1 1991, 6, 1608–1610. 52 Gagnon, R., Grogan, G., Levitt, M.S., Roberts, S.M., Wan, P.W.H., Willetts, A.J. Biological Baeyer–Villiger oxidation of some monocyclic and bicyclic ketones using monooxygenases from Acinetobacter calcoaceticus NCIMB 9871 and Pseudomonas putida NCIMB 10007. J Chem Soc, Perkin Trans 1 1994, 18, 2537–2543. 53 Vrtis, J.M., White, A.K., Metcalf, W.W., van der Donk, W.A. Phosphite dehydrogenase: a versatile cofactorregeneration enzyme. Angew Chem Int Ed 2002, 41, 3257–3259. 54 Loida, P.J., Sligar, S.G. Molecular recognition in cytochrome P-450: mechanism for the control of uncoupling reactions. Biochemistry 1993, 32, 11530–11538. 55 de Gonzalo, G., Torres Pazmino, D.E., Ottolina, G., Fraaije, M.W., Carrea, G. Oxidations catalyzed by phenylacetone monooxygenase from Thermobifi da fusca. Tetrahedron Asymmetry 2005, 16, 3077–3083. 56 Zambianchi, F., Pasta, P., Carrea, G., Colonna, S., Gaggero, N., Woodley, J.M. Use of isolated cyclohexanone

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monooxygenase from recombinant Escherichia coli as a biocatalyst for Baeyer–Villiger and sulfide oxidations. Biotechnol Bioeng 2002, 78, 489–496. Atia, K.S. Co-immobilization of cyclohexanone monooxygenase and glucose-6-phosphate dehydrogenase onto polyethylenimine-porous agarose polymeric composite using γ irradiation to use in biotechnological processes. Radic Phys Chem 2005, 73, 91–99. Walton, A.Z., Stewart, J.D. Understanding and improving NADPHdependent reactions by nongrowing Escherichia coli cells. Biotechnol Prog 2004, 20, 403–411. de Gonzalo, G., Ottolina, G., Zambianchi, F., Fraaije, M.W., Carrea, G. Biocatalytic properties of Baeyer– Villiger monooxygenases in aqueous– organic media. J Mol Catal, B Enzym 2006, 39, 91–97. Kamerbeek, N.M., Olsthoorn, A.J., Fraaije, M.W., Janssen, D.B. Substrate specificity and enantioselectivity of 4hydroxyacetophenone monooxygenase. Appl Environ Microbiol 2003, 69, 419–426. de Gonzalo, G., Torres Pazmino, D.E., Ottolina, G., Fraaije, M.W., Carrea, G. 4Hydroxyacetophenone monooxygenase from Pseudomonas fluorescens ACB as an oxidative biocatalyst in the synthesis of optically active sulfoxides. Tetrahedron Asymmetry 2006, 17, 130–135. Moonen, M.J.H., Westphal, A.H., Rietjens, I.M.C.M., van Berkel, W.J.H. Enzymatic Baeyer–Villiger oxidation of benzaldehydes. Adv Synth Catal 2005, 347, 1027–1034. Mihovilovic, M.D., Müller, B., Schultze, A., Stanetty, P., Kayser, M.M. An enantiodivergent trend in microbial Baeyer–Villiger oxidations of functionalized pentalenones by recombinant whole cells expressing monooxygenases from Acinetobacter and Pseudomonas. Eur J Org Chem 2003, 2243–2249. Wang, S., Kayser, M.M., Iwaki, H., Lau, P.C.K. Monooxygenase-catalyzed Baeyer–Villiger oxidations: CHMO

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4 The Bacterial Cytochrome P450 Monooxygenases: P450cam and P450BM-3 Vlada B. Urlacher, Stephen G. Bell, and Luet-Lok Wong

4.1 Introduction

Cytochrome P450 monooxygenases are versatile biocatalysts that insert an oxygen atom derived from atmospheric dioxygen into activated and non-activated C–H bonds in a vast range of molecules, often in a regio- and stereoselective manner. They play pivotal roles in primary and secondary metabolic pathways and in drug degradation. The number of identified CYP genes and enzymes is constantly increasing through studies on genomes and microbial screening (http://drnelson. utmem.edu/CytochromeP450.html). Numerous prokaryotic P450 enzymes have been identified and characterized in the past 20 years [1, 2]. Among them P450cam (CYP101A1) from Pseudomonas putida [3] and P450BM-3 (CYP102A1) from Bacillus megaterium [4, 5] are the best characterized because of their ease of heterologous expression and soluble nature. The structure, catalytic mechanism, and biochemistry of these two enzymes have been studied in detail [6]. The selective C—H bond oxidation activity of P450 enzymes under mild conditions using oxygen as the oxidizing agent has no equivalent in classical synthetic methodologies. In addition the alcohol products of substrate oxidation are more reactive to further chemical conversions. This is the basis of xenobiotic metabolism, detoxification and degradation. Hence P450 enzymes have significant potential applications in synthesis and bioremediation.

4.2 Biotransformation by Bacterial P450 Enzymes

The monooxygenase activity of P450 enzymes requires two electrons from NAD(P)H to activate oxygen for substrate oxidation: NAD(P)H + RH + O2 + H + → R–OH + NAD(P) + + H2O

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The two electrons are delivered to the P450 enzyme by electron transfer proteins which differ between organisms [6]. Most bacterial P450s utilize two electron transfer proteins, commonly a flavin-dependent reductase and an iron-sulfur ferredoxin. Mammalian systems use a diflavin protein, NADPH-cytochrome P450 reductase, as a common electron transfer shuttle, while some bacterial systems such as the CYP102 family and P450RhF are self-sufficient proteins with fused electron transfer and P450 monooxygenase domains [6]. Biotransformation applications of P450 enzymes require a high level of enzyme expression, fast substrate oxidation rate, system stability for high total turnover, and high selectivity of product formation. Bacterial P450 monooxygenases are soluble, more stable than their eukaryotic counterparts, and exhibit higher catalytic activities and expression rates in recombinant hosts. These properties make them from a practical standpoint, promising candidates for biocatalysis. However, they often have narrow substrate specificity and only moderate selectivity. Protein engineering and directed evolution can be used to broaden the substrate range as well as to increase the turnover activity, coupling, and product selectivity. Another important factor is the yield of product based on NAD(P)H consumed, or the coupling efficiency. The enzyme/substrate match for non-natural substrates is often suboptimal and results in reducing equivalents from NAD(P)H being channelled away from product formation to give instead hydrogen peroxide and water. These “uncoupling” pathways have to be minimized during enzyme development. The high cost of the NAD(P)H cofactor is a potential barrier to applying P450 enzymes in synthesis. A number of approaches to overcome this have been explored. Whole cell biotransformation faces the challenge of large reaction volume, slow mass transport, substrate and product toxicity, further oxidation or conversion of products by other cellular enzymes, and product recovery. The host organism can be immobilized to facilitate product recovery (e.g. in a flow reactor system), although mass transport will be slowed down further. Novel approaches have been developed to alter the property of the host cell wall to improve substrate uptake [7]. Two-phase reactions could minimize toxicity and product yields approaching 10 g L −1 have been achieved [8]. Applications in vitro require cofactor recycling; enzymatic regeneration has been successfully used for P450BM-3 utilizing D/L-isocitrate dehydrogenase [9], formate dehydrogenase [10], and alcohol dehydrogenase [11]. Other approaches are reviewed elsewhere in this volume (see Chapter 12). One attractive route that eliminates the need for cofactors is to supply electrons from an electrode [12], in particular if the enzyme is immobilized on the electrode to protect it from solvents and facilitate product recovery. However, this approach is still in early stage development. Turnover rates are slower than in solution and total turnover numbers are low. New avenues for delivering the electrons to fully functional enzymes at an electrode are required. The need for electrons could also be circumvented by the peroxide shunt in which alkyl peroxides and hydrogen peroxide or its precursors convert the ferric heme directly to the active ferryl intermediate. Enzyme inactivation and relatively slow reactions compared to the physiological activity are major obstacles. Directed evolution of P450BM-3 has generated mutants that

4.2 Biotransformation by Bacterial P450 Enzymes

are more active in the peroxide shunt and more stable to inactivation [13]. The bacterial enzymes, which work in an aqueous environment, are exposed to organics during in vitro oxidations. Immobilizing the enzymes could reduce the denaturing effect of solvents [14]. The resistance of P450BM-3 to organic solvents such as DMSO has been increased by directed evolution [15]. Numerous expression systems have been applied for the heterologous expression of the P450cam and P450BM-3 in E. coli for whole cell biotransformation. Expression of the single polypeptide enzyme P450BM-3 is more straightforward: examples include pUC13 in E. coli JM103 [16], pGLW11 (pKK223–3 derivative with a tac promoter) in DH5α [17], pCWOri+ (tandem tac promoters) in a catalase-deficient strain [18], and pCYTEXP1, which contains the tandem promoters PR and PL , in DH5α [19]. In the pET28a+ construct, with a T7 promoter, P450BM3 levels of up to 1200 nmol L −1 (∼140 mg L −1) were reported [10]. This system is appropriate for expression both in microplate format and fed-batch fermentation for high-level enzyme production. The first functional expression of the three-component P450cam system was in the form of a triple fusion (putidaredoxin reductase-putidaredoxin-P450cam) protein [20]. The activity of this in vivo system was lower than the in vitro reconstituted system because the linker peptides slowed down electron transfer. The pCWori+ vector was used to express the three proteins in a tricistronic system with each protein under the control of its own ribosome-binding site [21]. The expression level of the P450cam enzyme was lower than when it was expressed separately. Interestingly, when P450cam mutants were expressed in this system the host cells took on a blue coloration due to indigo formation via indole oxidation. The electron transfer proteins putidaredoxin reductase and putidaredoxin have been expressed from a pUC18 vector in the same host as the P450cam enzyme which was expressed from a pACYC177 derivative with the arabinose promoter to provide a system capable of oxidizing 1 g L −1 of camphor [22]. With increasing availability of metagenomes, genome sequences, and sophisticated heterologous expression methods, new P450 enzymes identified by bioinformatics methods are readily expressed and isolated. These new enzymes will greatly expand the range of compounds that can be oxidized but equally important will be the different product selectivity to existing enzymes. However, even if substrates and their novel biotransformation were established, a significant difficulty facing the application of new enzymes is the low stability and activity of electron transfer chains to support the enzymatic activity. Many P450 genes are found without the associated electron transfer proteins immediately upstream or downstream, or nearby in the genome sequence. It then becomes necessary to screen many candidate genes to reconstitute the activity. Known electron transfer proteins from other P450 systems (e.g. P450cam, P450lin, and P450cin [6]) can also be tested. However, such “cross” reactivities are often low. Notably mammalian P450 activity can be supported by the human and yeast NADPH-cytochrome P450 reductase. This difficulty with bacterial systems makes the self-sufficient P450 systems such as the CYP102 family and P450RhF very attractive because the electron transport chains in these enzymes are fused to the P450 domain.

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There is a real need for competent electron transfer chains for reconstituting the activity of the numerous other P450 enzymes that are “isolated” in the genome sequences but which display novel substrate specificity. The putidaredoxin reductase-putidaredoxin system of P450cam [23], the fused flavin-ferredoxin domain of P450RhF [24], and a ferredoxin reductase-ferredoxin system from a Mycobacterium strain [25], have been used with some success. However, more of such electron transfer systems are needed to cover the diverse P450 enzymes being discovered.

4.3 General Features of P450cam and P450BM-3

P450cam catalyzes the stereospecific oxidation of camphor to 5-exo-hydroxycamphor (Scheme 4.1). It was the first P450 enzyme to be structurally characterized [26, 27], identifying the amino acid side-chains that define the active site and contact the substrate. Camphor is bound by numerous non-covalent contacts and a hydrogen bond between the camphor carbonyl and the phenol side-chain of Y96, leading to a specific orientation with C5 directly above the heme iron (Fig. 4.1). Atkins and Sligar were the first to investigate the effects of active site

Fig. 4.1 The active structure of P450cam with bound camphor showing the residues that contact the substrate. The hydrogen bond between the camphor carbonyl and Y96 side-chain plays a crucial role in orientating the substrate for regiospecific oxidation at C5 which is located above the heme. Hydrogen binding between the proximal thiolate and amide NH of L358 modulates the heme redox potential.


changes on P450cam activity and substrate specificity by introducing the Y96F substitution to remove the hydrogen bond to camphor [28]. This mutation weakened camphor binding by a factor of 2, and other oxidation products (total 8%) were observed in addition to 5-exo-hydroxycamphor. Hence the main role of the hydrogen bond is not tight camphor binding but to orientate the substrate within the binding site and determine the regioselectivity of oxidation.

Scheme 4.1

The role of hydrophobic enzyme/substrate contacts was explored with norcamphor and 1-methylnorcamphor. For wild-type P450cam, the absence of methyl substituents in norcamphor compared to camphor lowered the selectivity to 45% of the 5-exo-hydroxy product, but the presence of just one methyl group in 1methylnorcamphor raised the selectivity to 82% [29]. The V295I mutation increased the selectivity for the 5-exo product for norcamphor oxidation; the larger Ile side-chain compensated for the two methyl groups missing in 1-methylnorcamphor compared with camphor. This was the first example of using active site mutations in P450 enzymes to compensate for structural differences between a target compound and the natural substrate. Montellano and co-workers modeled the binding of non-natural substrates by wild-type P450cam and the L244A mutant using a computer docking protocol. When the turnover rates of these substrates were subsequently determined they showed significant correlation with the modeling data [30, 31]. Nicotine binding and oxidation by wild-type P450cam has been investigated [32]. Computer modeling agreed with the product being from oxidation at the 5′ methylene group. However, the crystal structure showed nicotine N-coordination to the heme FeIII center in an unproductive binding orientation. The structure of the FeII(CO) complex, which could serve as a model for the FeII(O2 ) complex in the catalytic cycle, showed that the nicotine nitrogen was displaced from the heme iron and the resultant binding orientation was consistent with 5′ methylene oxidation [33]. This redox-induced switch might also operate in other P450 enzymes for substrates with a hetero-atom with an available lone pair of electrons. P450BM-3 (CYP102A1) is a fusion protein composed of the N-terminal P450 monooxygenase domain linked to the C-terminal diflavin reductase domain. This protein organization is responsible for the high activity of this enzyme compared with other known P450s. Recently other members of the CYP102A family have been characterized (e.g. CYP102A2 and CYP102A3 from Bacillus subtilis) [34]. All three enzymes catalyze hydroxylation of medium to long-chain saturated fatty acids (Scheme 4.2) as well as epoxidation of unsaturated fatty acids. The physiological role of these fatty acid hydroxylases in Bacillus strains is unclear, but their

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high activity and selectivity towards iso- and anteiso-fatty acids suggest a role in regulating membrane fluidity [35]. P450BM-3 and its mutant A74G/F87V/L188Q demonstrated activity (up to 1200 min−1) towards a range of highly branched fatty acids and gave only one product in each reaction [36]. Highly branched fatty acids have polyketide-like structures. Selective functionalization of such substances could offer convenient routes to biologically active compounds such as macrolide antibiotics.

Scheme 4.2

High-resolution X-ray crystal structures are available for substrate-free [37], palmitoleic acid-bound [38], and N-palmitoylglycine-bound [39] forms of P450BM3. Ligand binding has been studied by solution and solid-state NMR methods [40, 41]. The active site of P450BM-3 (Fig. 4.2) consists of a long hydrophobic channel, extending from the heme to the protein surface [38]. Analysis of crystal structures, followed by site-directed mutagenesis, revealed the important role of R47 and Y51 at the entrance of the substrate access channel. These two residues interact with the carboxylate moiety and are thus crucial for the proper positioning of fatty acid substrates. The guanidinium group of R47 plays a particularly prominent role [42, 43], while substitutions at Y51 appeared to have less impact [44]. The R47E, R47A, and R47G mutants retained their activity towards C12–C16 fatty

Fig. 4.2 The active site structure of P450BM-3 with palmitoleic acid (Palm) bound. The substrate carboxylate group interacts with R47 and Y51 at the entrance of the substrate channel while the aliphatic chain winds its way towards the heme. The F87 side-chain blocks access of the substrate to the heme iron and must move out of the way during the catalytic cycle to facilitate substrate oxidation.


acids, but the kcat/K m values were 5- to 15-fold lower than those of the wild type [43–45]. The highly conserved active site residue F87 is important for correct orientation of the fatty acid hydrocarbon chain. Comparison of substrate-free and substratebound crystal structures of P450BM-3 revealed a substantial conformational difference that is caused by the F87 phenyl ring [37, 38]. Mutations at F87 can affect the activity and selectivity [46]. An unfavorable substitution F87 could lead to irreversible conformational changes during catalytic turnover which resulted in a decrease or complete loss of catalytic competence [44]. 4.3.1 Aromatic Compounds

Wild-type P450cam has been shown to oxidize ethylbenzene to 1-phenylethanol with some stereoselectivity but low activity and coupling (Scheme 4.3) [47]. Substituting residues in the upper region of the substrate pocket (e.g. the T185L and V247M mutations, see Fig. 4.1) increased the coupling 2-fold, while similar mutations near the porphyrin plane (e.g. T101M and V295I) lowered the coupling efficiency. Bulky side-chains high in the pocket force substrates to bind closer to the heme and retard uncoupling. However, such substitutions close to the heme tend to increase uncoupling because there is less space in the vicinity of the heme and substrates may have to bind further away. The rate and coupling of alkylbenzene oxidation were increased by the T185L and T185F mutations but there was little correlation between side-chain properties and activity or product selectivity [48]. 2-Phenylethanol is the more desirable fine chemical product from ethylbenzene oxidation but this compound was not formed. The enzymes instead attacked the more reactive benzylic C–H bond to give 1-phenylethanol, suggesting that the substrate is sufficiently mobile within the active site to allow the enzyme to sample different parts of the substrate and the most reactive C–H bond is attacked.

Scheme 4.3

Styrene and naphthalene were oxidized by the Y96F mutant at rates 25 and 140 times those of wild-type P450cam, respectively (Scheme 4.4) [49, 50]. Both wildtype P450cam and the Y96F mutant oxidized tetralin stereospecifically to the 1-R alcohol (Scheme 4.4) [51, 52]. Styrene oxide, 1-naphthol and 1-tetralol are useful intermediates in synthesis. Both the NADH turnover rate and coupling were greatly improved for the Y96F mutant, demonstrating the value of increased active site hydrophobicity for the oxidation of hydrophobic molecules. The increased heme spin state shift induced by styrene binding to the Y96F mutant was accompanied by a more positive heme reduction potential [53]. The activity of this mutant was further increased by incorporating the V247L mutation to improve

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the enzyme–substrate match [21]. The Y96A mutation gave less dramatic increases because the larger active site resulted in substrate mobility and hence more uncoupling. However the Y96A mutant oxidized diphenylmethane, diphenylether and diphenylamine at the para position while the wild-type and Y96F mutant were inactive (Scheme 4.4) [54], demonstrating the utility of creating space in the active site for larger substrates. P450cam mutants with increased diphenylmethane oxidation activity over the wild-type enzyme have been identified by screening for colored indole oxidation products following site-saturation mutagenesis at Y96 and F98 [55]. These active mutants also contain mutations that increase the active site volume by replacing Y96 and F98 with residues that have smaller side-chains.

Scheme 4.4

The planarity and rigidity of polycyclic aromatic hydrocarbons (Scheme 4.5) impinge significantly on coupling efficiencies for their oxidation by P450cam [50]. For example, the NADH turnover rates for pyrene oxidation by the Y96A and Y96F mutants were 3 times those of wild type, although the couplings were generally low.

Scheme 4.5


Polychlorinated aromatics are hazardous environmental contaminants because of their lipid solubility, toxicity, and potential carcinogenicity. Many of these compounds are resistant to biodegradation, and the recalcitrance increases with the degree of chlorination. Chlorinated phenols, being more reactive, are more readily degraded. Biodegradation systems can therefore be generated by genetically augmenting chlorophenol-degrading organisms with P450 enzymes that oxidize polychlorinated benzenes to phenols (Scheme 4.6) [57]. P450cam was engineered to oxidize polychlorinated benzenes by using the Y96F mutation to promote hydrophobic molecule binding and the F87W, Y96W, and V247L mutations to force the benzenes closer to the heme. The mutants were up to three orders of magnitude more active than the wild-type enzyme for oxidizing di-, tri-, and tetrachlorobenzenes, with coupling efficiencies as high as 95%. The turnover activity and coupling for pentachlorobenzene (PeCB) oxidation were low [58].

Scheme 4.6

The crystal structure of the F87W/Y96F/V247L mutant complexed with 1,3,5trichlorobenzene (TCB) showed multiple substrate binding orientations with different angles between the benzene ring and the porphyrin [59]. The most productive orientation had TCB almost parallel to, and in van der Waals contact with, the heme (Fig. 4.3a). The structure showed that PeCB could not bind in this orientation because of steric interference with Leu244. The L244A mutation was therefore introduced and the F87W/Y96F/L244A/V247L mutant oxidized PeCB 45 times faster than the parent mutant, while hexachlorobenzene oxidation activity was increased 200-fold. Both compounds are oxidized to pentachlorophenol. The crystal structure of this mutant complexed with PeCB (Fig. 4.3b, unpublished results) strikingly confirmed that the substrate was bound in the parallel orientation, with a chlorine being accommodated in the space created by the L244A mutation. Introduction of a gene cassette encoding this mutant and the electron transfer proteins putidaredoxin reductase and putidaredoxin into a pentachlorophenol-degrading Sphingobium strain generated a novel microorganism capable of degrading hexachlorobenzene [60]. Substitutions at F87 in P450BM-3 have been used to alter the side-chain volume at this residue and increase the activity and alter the enantioselectivity of propylbenzene (Scheme 4.7) and 3-chlorostyrene oxidation. In the case of 3-chlorostyrene the F87G mutant gave (R)-chlorostyrene oxide with up to 96% ee [61]. Experiments with styrene revealed that other positions in and outside of the binding site also affected the absolute configuration of the product and could even

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Fig. 4.3 The active site structures of (a) the 1,3,5-TCB complex of the F87W/Y96F/V247L mutant of P450cam and (b) the PeCB complex of the F87W/Y96F/L244A/V247L mutant. Both compounds are bound with the aromatic ring almost parallel to the heme porphyrin. The structure of 1,3,5-TCB

complex shows that the L244 side-chain will sterically hinder the binding of PeCB. Introduction of the L244A mutation generates space to accommodate the extra chlorines in PeCB and results in a 45-fold increase in PeCB oxidation activity.


lead to inversion of the enantioselectivity. The enantiomeric excess of the reaction product of styrene oxidation ranged from 58% ee (S)-styrene oxide (A74E/F87V/P386S) via 49% (R)-styrene oxide (F87A) and 65% (R)-styrene oxide (A74G/F87V/L188Q) to 92% (R)-styrene oxide for the F87G mutant (Scheme 4.7) [62].

Scheme 4.7

The A74G/F87V/L188Q mutant oxidized naphthalene and three-ring polyaromatics such as fluorene and acenaphthene three orders of magnitude faster than the wild-type enzyme [63]. A different approach combined the hydrophobic R47L/ Y51F mutations at the entrance to the substrate channel with the F87A, I263A, and A264G mutations in the active site to give similar orders of magnitude increases in activity for oxidizing polyaromatics with up to four rings [17]. The A74G/F87V/L188Q mutant showed faster NADPH turnover but the R47/Y51F series of mutants had higher couplings, such that the overall substrate oxidation activities were comparable. Polychlorinated aromatic oxidation by CYP102 enzymes has been less studied, but the A74G/F87V/L188Q mutant has been shown to oxidize polychlorinated dioxins [64]. The activity pattern was similar to that for chlorinated benzene oxidation by P450cam (i.e. dioxins with up to three chlorines were oxidized but 2,3,7,8-tetrachlorodibenzodioxin (TCDD) was not oxidized). The enzyme attacked at unsubstituted positions and NIH shifts involving chlorines were observed. 4.3.2 Alkanes and Alicyclics

Engineering P450 enzymes into an alkane hydroxylase is an area of ongoing interest. A methane-oxidizing P450 enzyme may have important applications in the energy and chemicals sector while terminal oxidation of medium and long-chain alkanes and alkenes leads to fine chemicals. The CYP102 family of medium-chain fatty acid hydroxylases is a natural choice for engineering for alkane oxidation. Wild-type P450BM-3 showed low activity for octane oxidation (80 min−1). The A74G/F87V/L188Q mutant oxidized octane with fast NADPH

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turnover (1760 min−1) and 40% coupling mainly to 3- and 4-octanol, with 17% 2octanol [65]. Almost all P450BM-3 mutants reported to date produce secondary alcohols. Only Peters et al. have reported a variant containing 15 mutations that produces 10% 1-octanol [66]. Wild-type CYP102A3 shares 64% homology to the monooxygenase domain of P450BM-3 and shows limited regioselectivity for octane oxidation. However, CYP102A3 has a more hydrophobic and narrower binding site than P450BM-3 and higher activity towards octane (110 min−1). The regioselectivity of CYP102A3 was changed by combination of error-prone PCR and site-directed mutagenesis to hydroxylate substrates not only at different subterminal carbons but also to a high extent at the terminal carbon. To enable high-throughput screening, a specific assay was developed, based on yeast alcohol dehydrogenase that is capable of discriminating between products of terminal and subterminal hydroxylation. Two CYP102A3 mutants, F88V/S189Q/A328V and S189Q/A328V, were identified and these produced 17% and 50% 1-octanol, respectively [67]. All three mutations occur in close proximity to the heme center and hinder approach of the substrate molecule to the heme, leading to terminal oxidation but which might also cause the poor coupling efficiencies of 5–7%. By screening for activity with a p-nitrophenoxyoctane oxidation assay, a derivative of a p-nitrophenoxyoctanoate oxidation assay [19], P450BM-3 was taken through multiple rounds of directed evolution to identify the mutant “139–3” which oxidized octane with high turnover rates and 66% selectivity for 2-octanol [68]. This mutant was also capable of oxidizing alkanes as short as butane and propane. Further rounds of directed evolution combined with designed mutations in the active site lead to activity improvements and significant changes in regioand stereoselectivity [66]. A mutant labeled “1–12G” yielded 82% 2-octanol, and several variants yielded small quantities of terminal hydroxylation products for a range of alkanes. An exciting development was the mutant 35-E11 prepared from the 9–10A by directed evolution while deliberately constricting the active site [69]. This mutant functions as an ethane hydroxylase with an ethanol formation rate of 0.4 min−1 and 0.8% coupling. High selectivity was observed in the epoxidation of terminal alkenes using a rationally designed variant of mutant 9–10A involving active site substitutions such as F87V and I263A [11]. P450cam has been redesigned for alkane oxidation. The size of the substrate and its shape in relation to the active site were crucial considerations. The alkane oxidation activity of the wild-type and Y96F mutants was highest for pentane/hexane and decreased steadily to octane [70]. Smaller substrate pockets generated by bulky substitutions such as V247L favored a sterically less demanding substrate such as hexane over 3-methylpentane [71]. Increases in activity towards smaller alkanes were achieved by introducing bulky residues high up in the active site to oblige the substrate to bind closer to the heme iron. The F87W/Y96F/T101L/ V247L mutant oxidized butane with a turnover rate of 750 min−1 compared with 0.4 min−1 for the wild-type enzyme [72]. Further bulky substitutions, together with the L358P mutation first used by Morishima and co-workers, were also added. The L358P mutation tightened the active site, pushed the heme towards the substrate,


and increased the donor strength of the proximal cysteine thiolate by eliminating a hydrogen bond between Leu358 and Cys357 which promoted O–O bond cleavage and reduced uncoupling [73, 74]. The F87W/Y96F/T101L/T185M/L244M/V247L/ G248A/L358P mutant oxidized propane at 500 min−1 with 86% coupling. The NADH turnover rate for ethane was even higher (∼800 min−1) but the coupling for ethanol formation was low (10.5%) [75]. Enantioselective hydroxylation of achiral substrates to chiral alcohols is an attractive route to fine chemicals and synthetic intermediates. P450cam was engineered to oxidize phenylcyclohexane selectively at C2, C3, or C4 of the cyclohexane ring [54, 76–78]. The Y96F mutant gave 81% cis-3-phenylcyclohexanol with 34% ee and the Y96F/V247A mutant gave 97% with 42% ee, while the Y96F/V247L mutant gave 83% trans-4-phenylcyclohexanol. Oxidation of a cyclopentanecarboxylic acid derivative by P450BM-3 mutants has been reported. The 139–3 mutant gave principally the S,S product while 1–12G formed the R,R diastereoisomer [79]. 2-Ethylhexanol is oxidized to 2-ethylhexanoic acid by P450cam [80, 81]. The F87W and Y96W mutants gave the acid exclusively, while the T185F mutation improved coupling but was less selective. The crystal structure of the wild type with the hexanol substrate bound revealed that the R-isomer was bound in a more ordered state and was the preferred substrate over the S-isomer. 4.3.3 Terpenoid Compounds

Terpenoid compounds constitute one of the largest groups of biological molecules. The parent terpene hydrocarbons are often readily available and their oxygenation gives derivatives which are fragrance and flavoring compounds, insect pheromones, and precursors to pharmaceuticals. The monoterpene (+)-α-pinene is structurally related to camphor. The Y96F mutant of P450cam showed strengthened monoterpene binding and increased oxidation rates and coupling [82]. Pinene was oxidized mainly to verbenol and pinene oxide while limonene gave mainly isopiperitenol and some limonene oxide. The proportion of epoxides was higher for mutations that increased the active site volume, indicating that substrate mobility would result in the reactive olefinic bond being referentially attacked. The F87W and V247L mutations were introduced as a means of reducing substrate mobility. Despite weakening substrate binding, both mutations substantially increased pinene oxidation rates and improved coupling efficiency while enhancing the selectivity for verbenol formation (70% for the F87W/Y96F/V247L mutant). The limonene oxidation rate of the F87W/Y96F/V247L mutant was lower than those for the F87W/Y96F and Y96F/V247L mutants but it showed substantially higher selectivity (∼90%) for isopiperitenol formation. The crystal structure of the triple mutant complexed with (+)-α-pinene revealed two substrate binding orientations. In one orientation C3 was positioned over the heme iron (Fig. 4.4a), consistent with verbenol formation. In the other the heme iron was closest to the allylic C10 methyl group, a configuration that should give myrtenol

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Fig. 4.4 The active site structure of the F87W/Y96F/V247L mutant of P450cam with showing the two binding orientations observed for (+)-α-pinene. The orientation in (a) is related to the camphor binding orientation in the wild type, with the bridgehead methyl group interacting with V295. The C3 allylic carbon is closest to the heme iron and (+)-verbenol is the predicted

product. The orientation in (b) should give rise to (+)-myrtenol but this is not observed in the products. Rapid interconversion between the two orientations results in the more reactive C—H bond at C3 to be preferentially attacked. There are subtle differences in the L244 and V247 side-chain conformations between the two orientations.


(Fig. 4.4b). Since no myrtenol was detected in practice, it was proposed that the two orientations interconvert, leading predominantly to oxidation of the allylic CH2 group at C3, which is significantly more reactive than the methyl group at C10. Analysis of enzyme/substrate interactions indicated that the F87W and V247L mutations had opposing effects on pinene orientation within the active site, and that the L244 side-chain contacted multiple substrate carbons. The F87W/Y96F/L244A mutant gave 86% (+)-cis-verbenol and the Y96F/L244A/ V247L mutant gave similar selectivity for C3 oxidation but 32% of (+)-verbenone was formed with 55% (+)-cis-verbenol (Scheme 4.8) [83]. Both verbenol and verbenone are active pheromones against various beetle species.

Scheme 4.8

(+)-Valencene, a sesquiterpene found in orange oil, is the likely biological precursor of (+)-nootkatone, a fragrance found in grapefruit juice. Valencene is larger and more hydrophobic than camphor. No valencene oxidation activity was found for wild type P450cam or the Y96A and Y96F mutants even though the Y96A mutant oxidized diphenylmethane which is of comparable size [84]. The total active site volume in the Y96A might be sufficient for valencene, but the topology was incorrect. The Y96F mutation was used to render the active site more hydrophobic, and smaller amino acids were introduced at F87 to alter the topology. Ideally the space created at the 87 side-chain should accommodate the isopropenyl group of valencene, which would place C2 closest to the heme iron for oxidation to nootkatol and thence to nootkatone. The 87/96 double mutants oxidized valencene at C2 and the effect of side-chain volume at 87 was subtle. The F87V mutation favored the oxidation of nootkatol but not of nootkatone while the F87A and F87L mutations both retarded the oxidation of nootkatol to nootkatone. Additional substitutions were introduced with a view to improving selectivity. The V247L mutation retarded the further oxidation of nootkatol while the L244A mutation had the opposite effect, promoting the oxidation of nootkatol and nootkatone. In combination studies the F87A/Y96F/L244A/V247L mutant gave 86% (+)-trans-nootkatol, 4% nootkatone and 10% of products from nootkatone oxidation (mainly 9-hydroxynootkatone) (Scheme 4.9) while the F87V/Y96F/L244A mutant gave 38% nootkatol and 47% nootkatone.

Scheme 4.9

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(+)-Valencene oxidation by a series of R47L/Y51F mutants of P450BM-3 has been investigated [84]. Compared with the P450cam mutants, the turnover activities of the P450BM-3 mutants were higher but they were much less selective. Nootkatol and nootkatone were minor products in some mutants but most gave many products. Moreover, nootkatone was oxidized more rapidly than valencene and hence the steady state concentration of nootkatone was low. The R47L/Y51F series of P450BM-3 mutants also oxidized pinene and limonene with higher activity than P450cam mutants but again the selectivity was low (Sowden and Wong, unpublished results). However, P450BM-3 mutants showed excellent regioselectivity for the oxidation of β-ionone (a sesquiterpenoids analog) to 4-hydroxy-βionone, with stereoselectivity of up to 39% ee for the R-isomer (Scheme 4.10) [85]. Such ionone oxidation products are flavor components and synthetic intermediates. The F87V mutant was 100 times more active than the wild type, and addition of the R47L/Y51F combination increased the activity 3-fold further. Screening after two rounds of random mutagenesis of the F87V mutant identified the triple mutant A74E/F87V/P386S which was also 300 times more active than the wild type. P386 is located on the surface of the enzyme molecule. Nevertheless, its substitution by serine dramatically increased the NADPH turnover rate during β-ionone hydroxylation. The crystal structure of the complex between the heme and FMN-containing domains of P450BM-3 suggested that the region between P382 and E387 might affect electron transfer from the FMN to the heme iron [86]. Therefore, mutations of P386 could influence the interaction between the monooxygenase domain and an FMN-binding reductase domain. Also remarkable was the effect of the substitution A74E which apparently narrowed the substrate channel and provided a better orientation of the substrate in the binding pocket.

Scheme 4.10

4.3.4 Human Metabolites

An extraordinarily broad substrate range and very diverse reactions catalyzed by eukaryotic P450s make them attractive for pharmaceutical research. However, the low stability and activity of these enzymes hinder their application for drug synthesis. Implementation of more stable and active bacterial P450s, which are able to accept substrates of the mammalian counterparts, would open up a route to the production of new pharmaceuticals.


Wild-type P450BM-3 (fatty acid hydroxylase) does not metabolize any drug-like compounds, but it can be tuned by protein design and directed evolution into an enzyme which oxidizes compounds typical for human P450 monooxygenases. The R47L/F87V/L188Q mutant was shown to metabolize testosterone (Scheme 4.11), amodiaquine, dextromethorphan, acetaminophen, and 3,4methylenedioxymethylamphetamine [87]. Reactions catalyzed by this mutant include not only hydroxylation, as in the case of testosterone, but also N-dealkylation of e.g. dextromethorphan, O-demethylenation of 3,4-methylenedioxymethylamphetamine, and dehydrogenation of acetaminophen. In general the activity of R47L/F87V/L188Q for these compounds was very low, but in the case of 3,4methylenedioxymethylamphetamine and acetaminophen it could be stimulated up to 70-fold by the addition of caffeine, a known activator of rat P450 3A2.

Scheme 4.11

Several mutants of P450BM-3 obtained by directed evolution have been shown to oxidize propranolol (Scheme 4.12), a multi-function β-adrenergic blocker. A mutant “9C1,” which has 14 amino acid substitutions, produced a very similar product pattern in a bioconversion of propranolol to the human CYP2D6 and CYP1A2 [88]. After further optimization by saturation mutagenesis several new mutants were identified, yielding different ratios of products. Among the produced metabolites, the ring-hydroxylated products are of particular importance as they have been demonstrated to be equipotent to propranolol as a β-receptor antagonist. Most of the substituted amino acids (R47, A74, A82, F87) of importance are located in the active site of the enzyme and had already been shown to affect the substrate range and selectivity.

Scheme 4.12

The mutant 9–10A/F87A, constructed on the basis of the 9–10A evolved previously for alkane hydroxylation, was able to hydroxylate the α-position of the peptide group of buspirone. Buspirone (Scheme 4.13) as well as its human metabolite, 6-hydroxybuspirone, are anti-anxiety agents. Remarkable is the high enantio- and regioselectivity of the enzyme in this reaction. (R)-6-Hydroxybuspirone was the sole product formed with > 99.5% ee [89].

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Scheme 4.13

4.4 The Scope of P450 Engineering

It is not surprising that the engineering of bacterial P450 systems has been focused on the best characterized systems, P450cam and P450BM-3. There have been significant successes from structure-based redesign, site saturation mutagenesis, and directed evolution by error-prone PCR as well as chimera construction [90–92]. All these methods have limitations but the results to date have been very promising. Furthermore, these approaches are increasingly being combined to further enhance activity and selectivity. P450cam and P450BM-3 have been engineered to oxidize large molecules such as pyrene, small molecules such as propane and ethane, highly branched fatty acids, as well as complex molecules such as terpenes, alicyclic compounds and pharmaceuticals. In general, it is relatively straightforward to obtain fast NAD(P)H turnover rates but the coupling yield and especially selectivity of product formation are far more challenging. P450BM-3 in particular appears to be finely balanced between the slow reacting substrate-free form and fast electron transfer upon structural perturbations induced by mutations. For instance, the R47L/Y51F/F87A, A74G/ F87V/L188Q, 1–12G and 9–10A mutants all show fast NADPH turnover rates with a range of substrates. However, in many cases the reactions are uncoupled and non-selective. Refinement of screening methods (e.g. by the use of alcohol dehydrogenases that can discriminate between primary and secondary alcohols) will play an increasingly important role. Recent directed evolution of the CYP102 family of enzymes has imparted terminal alkane oxidation activity to these enzymes which naturally target subterminal positions [66, 67]. Similarly P450cam has been engineered for fast limonene oxidation but perillyl alcohol, a desirable terpenoid that arises from oxidation of a methyl substituent, was not observed (Scheme 4.14). Instead products from oxidation at the more reactive olefinic double bond and allylic CH2 groups dominated [82]. On the other hand the CYP153 family of enzymes are highly selective for terminal C–H bond oxidation, converting octane to 1-octanol and limonene to perillyl alcohol with almost total selectivity [8]. Therefore the discovery of novel activity by isolation and characterization of new P450 enzymes from diverse organisms still has a crucial role to play, together with protein engineering, for developing P450 biooxidation applications.

References

Scheme 4.14

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62 Eiben, S., Kaysser, L., Maurer, S., Kuhnel, K., Urlacher, V.B., Schmid, R.D. Preparative use of isolated CYP102 monooxygenases – A critical appraisal. J Biotechnol 2006, 124, 662–669. 63 Li, Q.S., Ogawa, J., Schmid, R.D., Shimizu, S. Engineering cytochrome P450 BM-3 for oxidation of polycyclic aromatic hydrocarbons. Appl Environ Microbiol 2001, 67, 5735. 64 Sulistyaningdyah, W.T., Ogawa, J., Li, Q.S., Shinkyo, R., Sakaki, T., Inouye, K., Schmid, R.D., Shimizu, S. Metabolism of polychlorinated dibenzo-p-dioxins by cytochrome P450 BM-3 and its mutant. Biotechnol Lett 2004, 26, 1857. 65 Appel, D., Lutz-Wahl, S., Fischer, P., Schwaneberg, U., Schmid, R.D. A P450 BM-3 mutant hydroxylates alkanes, cycloalkanes, arenes and heteroarenes. J Biotechnol 2001, 88, 167. 66 Peters, M.W., Meinhold, P., Glieder, A., Arnold, F.H. Regio- and enantioselective alkane hydroxylation with engineered cytochromes P450 BM-3. J Am Chem Soc 2003, 125, 13442. 67 Lentz, O., Feenstra, A., Habicher, T., Hauer, B., Schmid, R.D., Urlacher, V.B. Altering the regioselectivity of cytochrome P450 CYP102A3 of Bacillus subtilis by using a new versatile assay system. Chembiochem 2006, 7, 345. 68 Glieder, A., Farinas, E.T., Arnold, F.H. Laboratory evolution of a soluble, selfsufficient, highly active alkane hydroxylase. Nat Biotechnol 2002, 20, 1135. 69 Meinhold, P., Peters, M.W., Chen, M.M., Takahashi, K., Arnold, F.H. Direct Conversion of ethane to ethanol by engineered cytochrome P450 BM3. Chembiochem 2005, 6, 1. 70 Stevenson, J.-A., Westlake, A.C.G., Whittock, C., Wong, L.-L. The catalytic oxidation of linear and branched alkanes by cytochrome P450cam. J Am Chem Soc 1996, 118, 12846. 71 Stevenson, J.-A., Bearpark, J.K., Wong, L.L. Engineering molecular recognition in alkane oxidation catalysed by cytochrome P450 cam. New J Chem 1998, 551.

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92 Otey, C.R., Landwehr, M., Endelman, J.B., Hiraga, K., Bloom, J.D., Arnold, F.H. Structure-guided recombination creates an artificial family of cytochromes P450. PLoS Biol 2006, 4, e112.

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5 Cytochrome P450 Redox Partner Systems: Biodiversity and Biotechnological Implications Andrew W. Munro, Hazel M. Girvan, Joseph P. McVey, and Kirsty J. McLean

5.1 Introduction

The cytochromes P450 (P450s) are an ever-growing family of heme b-containing monooxygenase enzymes found in all domains of life [1]. The vast majority of this enzyme superfamily catalyze the reductive scission of dioxygen, which is bound to the heme iron. The process requires the consecutive delivery of two electrons to the heme iron, usually mediated by interaction with redox partner enzymes containing flavin or iron–sulfur centers [2]. Ultimately these electrons are almost invariably derived from the nicotinamide coenzymes NADH or NADPH. In order that the potential of P450 enzymes for biotechnologically relevant transformations can be realized, a natural and/or efficient redox partner system must be defined and conditions enabling well-coupled electron transfer and substrate oxygenation established. The identification of appropriate P450 redox partners thus brings an additional level of complexity into the system, and the successful exploitation of P450s requires a detailed understanding of mechanism and (where possible) structure of the relevant P450 and its redox partner(s). The last 10–15 years has seen a dramatic change in our understanding of P450 redox partners, their varied nature, structures and mechanisms. This chapter addresses our current state of knowledge of P450 redox systems and their biological diversity, and also considers the application of different strategies (including chemical and electrode systems) for driving P450 oxygenase systems and enhancing their applications in industrial biotransformations.

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5 Cytochrome P450 Redox Partner Systems: Biodiversity and Biotechnological Implications

5.2 P450 Redox Partners 5.2.1 A “Historical” Perspective

The P450s were recognized and defined as a distinct class of hemoproteins only ∼50 years ago. The enzymes form reduced (ferrous) iron–carbon monoxide adducts in which the major heme absorption band (the Soret band) shifts to ∼450 nm for the native form of the enzyme [3, 4]. Pivotal to this spectral characteristic is the axial coordination of the heme iron by a cysteine thiolate in all the P450s. The phylogenetically conserved cysteinate is termed the proximal ligand to the heme iron, with the distal ligand generally considered to be a weakly bound water molecule [5]. In early studies of P450 enzymes, two types of model system were characterized in detail. The first of these were membranous P450s derived from mammalian liver microsomes, which source their electrons from the diflavin (FAD- and FMN-containing enzyme NADPH-cytochrome P450 reductase (CPR) [6] (Fig. 5.1a). The second model system characterized from the 1960s onwards was the Pseudomonas putida camphor hydroxylase P450cam (now formally classified in the P450 gene superfamily as CYP101A1) [7]. P450cam participates in a catabolic pathway that facilitates the breakdown of camphor in the pseudomonad, and its use as a sole source of carbon for energetic requirements [8]. P450cam obtains its electrons from the iron–sulfur (2Fe-2S cluster-containing) protein putidaredoxin (Pd), which in turn sources electrons from the FAD-containing, NADH-dependent protein putidaredoxin reductase (PdR) [9, 10] (Fig. 5.1b). These two types of apparatus have come to be known as class I (CPR) and class II (PdR/Pd-type) P450 reductase systems, although (as discussed in more detail

Fig. 5.1 Protein components of class I and class II P450 redox systems. (a) Membraneassociated components of a class II P450 redox system. These are represented by the structure of mammalian CYP2C5 (PDB code 1DT6) and that of rat cytochrome P450 reductase (1AMO) [48, 146]. (b) The three components of the best-characterized class I P450 redox systems – Pseudomonas putida

P450cam. These are (from left) the P450 itself (CYP101A1, 1DZ6), the 2Fe-2S clustercontaining putidaredoxin (1PDX) and the FAD-binding putidaredoxin reductase (1Q1R) [5, 9, 10]. Helical and sheet segments of the proteins are distinctly colored. Bound cofactors are shown in spacefill representation: heme (red), FMN (orange), FAD (yellow) and 2Fe-2S (orange-blue)

5.2 P450 Redox Partners

below) far more diverse types of P450 redox systems have been identified in recent years. 5.2.2 The P450 Catalytic Cycle and Electron Transfer Events

In the classical P450 catalytic cycle (Fig. 5.2), the delivery of the fi rst electron to the P450 heme iron reduces it from the ferric (FeIII) to the ferrous (FeII) form. The FeII binds dioxygen, converting to a ferrous-dioxy complex. The timely delivery of the second electron converts this species to a ferric peroxy form. Thereafter, the species is protonated to form a ferric hydroperoxy intermediate, and then further protonated to form a high valent iron–oxo complex (compound I) with release of a molecule of water through heterolytic cleavage of the dioxygen bond in the preceding intermediate. Compound I is considered to be the intermediate catalyzing the majority of P450 reactions, although the ferric hydroperoxy intermediate may also participate in some P450-dependent catalytic reactions [11]. Since most P450 redox systems use electrons derived from the oxidation of the coenzymes NAD(P)H, the two electrons required for P450 oxidations should be provided to the redox partner as a hydride ion from the coenzyme. However, as is clear in the cycle shown in Fig. 5.2, these electrons are then required to be delivered singly at distinct points in the catalytic cycle – either side of the binding of dioxygen to the heme iron. Thus, the role of the redox partner systems is to take two electrons (as hydride ion) from NAD(P)H and to deliver these one at a time to the P450. The first electron is donated to the FeIII form, reducing it to FeII. In systems such as P450cam and the Bacillus megaterium flavocytochrome P450BM-3 (CYP102A1, see below), there is clear regulation over the transfer of electrons to the ferric heme iron mediated by the binding of substrate molecules (camphor and its analogs for P450cam, fatty acids for P450BM-3) [12, 13]. For these “well-regulated” P450s, binding of substrate in the active site induces the dissociation of a water molecule that is bound relatively weakly as the 6th coordinating ligand (trans to the thiolate) to the heme iron. This, in turn, induces an equilibrium shift of the heme iron spin-state (from low-spin S = 1/2 to high-spin S = 5/2) and a positive shift in heme iron reduction potential of the order of 130–140 mV [12, 13]. The increased potential brings the heme iron into range for efficient electron transfer from the redox partner (Pd for P450cam, a fused CPR for P450BM-3), and ensures that rapid electron transfer to the heme iron occurs only when substrate is available for oxygenation [14]. Similar substrate-dependent heme iron dehydration and redox potential shifts are seen in other bacterial P450 systems [e.g. 15, 16], and substrate-induced heme iron spin-state shift is also observed widely in eukaryotic P450 systems [e.g. 17]. However binding of substrate to P450s is not always a prerequisite for electron transfer from the redox partner, and this brings with it the likelihood of wastage of the reducing equivalents in the non-specific reduction of bound dioxygen – so called “uncoupling” of the P450 catalytic cycle (see Fig. 5.2 and the more detailed discussion below) [18].

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Fig. 5.2 Catalytic cycle of cytochrome P450. The resting state of a P450 is the ferric (FeIII) form, usually with water present as the distal ligand to the heme iron. In the first step, the water is displaced by binding of a substrate (R-H). The first electron (e−) transfer from the redox partner reduces the complex to the ferrous (FeII) form, which can then bind dioxygen (O2). The ferrous-oxy complex formed may also be presented as the isoelectronic ferric-superoxy form. The delivery of a second electron converts this to a ferric peroxy species, which is then protonated to the ferric hydroperoxy state, and then to compound I (presented as a ferryl-oxo porphyrin radical cation) with loss of water. Compound I oxygenates the bound

substrate (to R-OH) and product dissociation leads to restoration of the ferric state and reassociation of water as the distal ligand to complete the catalytic cycle. Collapse of unstable oxy intermediates leads to uncoupling of the cycle and reformation of the ferric form of P450. The ferrous oxy complex dissociates superoxide, while the ferric hydroperoxy complex collapses with peroxide generation. The reaction can be forced in the productive direction by addition of hydrogen peroxide (or an organic peroxide) to resting substrate-bound P450 (the so-called “peroxide shunt” pathway). Collapse of compound I occurs with generation of water [18].

Following delivery of the first electron, dioxygen binds the ferrous iron, and then the second electron should be delivered to avoid collapse of the ferrous-oxy species back to the ferric resting state of the iron – with wasteful production of superoxide radical. Subsequent steps in the P450 cycle are considered to be relatively fast with respect to the electron transfer events, and as a result the later


transient intermediates have proven difficult to characterize spectroscopically or structurally. Compelling evidence is available for the formation of the ferric hydroperoxy intermediate, but compound I (the ferryl oxo species following immediately from ferric hydroperoxy in the catalytic cycle) has proven much more elusive [e.g. 19]. Thus, the P450 redox partners have the task of “splitting” the two electron batch derived from nicotinamide coenzyme and passing single electrons to the P450 at discrete points in the cycle. To do this, the redox partner systems usually have more than one redox active cofactor, and use the first acceptor cofactor (a flavin) to accept the hydride ion from NAD(P)H and the second donor cofactor (usually another flavin or an iron–sulfur cluster) to transport single electrons between the donor flavin and the heme iron. The composition of the major P450 redox systems is discussed below. 5.2.3 P450cam and its Reductase System

The most intensively studied class I P450 redox system is P450cam along with its PdR and Pd redox partners. All components of the system are soluble proteins and interact with one another in the cytosol of Pseudomonas putida. The atomic structures of all three components of the system have been determined, and the P450 component has been structurally characterized in a number of different substrate- and ligand-bound forms [e.g. 9, 10, 20, 21]. PdR is a 48 kDa FAD-binding protein which is reduced by NADH. Pd is a 11 kDa 2Fe-2S cluster-containing ferredoxin (Fig. 5.1b). The system couples NADH oxidation tightly to the formation of 5-exo-hydroxycamphor, with Pd acting as a single electron carrier between the reductase and the P450. Substrate (camphor) binding to P450cam results in modulation of the heme iron spin-state equilibrium, with the P450 converting from a predominantly low-spin ferric form to a predominantly high-spin form at substrate saturation [22]. A large positive shift in heme iron reduction potential (∼130 mV from ca. −300 mV to −170 mV) accompanies this conversion, acting as a thermodynamic trigger for electron transfer from Pd to the substrate-bound P450 [23]. The steady-state turnover of P450cam in camphor hydroxylation is rate-limited by electron transfer from (but not binding of) Pd. At high Pd concentrations the first electron transfer reaction to ferric P450cam is rate-limiting (measured at ∼15 s−1 at 4 ºC), but at more physiologically relevant Pd concentrations the second electron transfer to the P450 becomes rate-limiting [8, 24]. Chemical modification studies of Pd indicated that a cluster of acidic residues is involved in interactions critical for electron transfer from PdR. Thus electrostatic binding is considered as important for productive PdR/Pd interactions [25]. The acidic cluster is in the same protein region as that recognized to be important for binding of the mammalian adrenal gland mitochondrial 2Fe-2S ferredoxin adrenodoxin to its reductase partner adrenodoxin reductase [26]. Cytochrome b5, a known effector (and likely second electron donor) for certain mammalian steroidogenic P450s, was

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found to bind tightly to P450cam, with binding mediated by electrostatic interactions between basic residues on the proximal heme face of the P450 and acidic residues on the b5 [27–29]. Pd competitively inhibits binding of b5 to P450cam, and this is consistent with a binding site for the redox partner close to the heme of the P450, and compatible with efficient electron transfer between the proteins [27, 30]. However, it is not clear to what extent the b5/Pd sites on P450cam overlap. What is clear is that a tryptophan residue at the C-terminus of Pd (Trp 106) is needed for efficient electron transfer to P450cam. However, removal of this residue leads to diminished binding (higher K d value) of Pd to P450cam, and there is no proof that Trp 106 participates directly in the electron transfer process [31–33]. Cytochrome P450cam (formally CYP101A1) was the first P450 enzyme to have its atomic structure resolved, and this structure has provided an important template on which several other P450 structures have been modeled over the years. P450cam has also been structurally resolved in several different substrate/ligand complexes and also, for example, in its reduced/CO-bound complex [e.g. 34]. Only in recent years have the atomic structures of Pd and PdR been resolved from protein crystals [9, 10]. The structure of PdR reveals a fold similar to those of disulfide reductases, and that of Pd is consistent with a key role for the C-terminal residue Trp106 in regulation of electron transfer to P450cam (and possibly from PdR). Structural studies of reduced Pd indicate that conformational changes occur in the protein that likely underlie the marked alterations in affinity previously observed for binding of the oxidized and reduced forms of Pd to P450cam [35]. Modeling of the interactions between PdR and Pd indicate that there is steric complementarity between the two proteins and hint at the possibility of multiple electron transfer pathways between the redox partners [36]. 5.2.4 Adrenodoxin and Adrenodoxin Reductase

The PdR/Pd system has a counterpart in the mammalian adrenal mitochondria, where the NADPH-dependent, FAD-containing adrenodoxin reductase (AdR) transfers electrons to the 2Fe-2S ferredoxin adrenodoxin (Adx), which in turn reduces mitochondrial P450 enzymes involved in steroidogenesis. Detailed studies have been done on the interactions between Adx and the mammalian CYP11A1 and CYP11B enzymes, involved in cholesterol side-chain cleavage (the first step in the steroid synthesis pathway) and in formation of cortisol and aldosterone, respectively [37]. The AdR and P450 proteins are membrane-associated, but the Ad is a soluble protein. The nature of this type of P450 redox system (analogous to that seen for P450cam and other bacterial P450s, but markedly different from that supporting P450s in other mammalian tissues) is consistent with the endosymbiont theory of mitochondrial evolution. Putidaredoxin shares ∼32 % amino acid sequence identity with bovine Adx, but Adx is not reduced efficiently by PdR, indicating important structural differences


required for interactions within the distinct redox systems [8, 25]. The bovine Adx is produced as a 186 amino acid precursor protein, which is then proteolytically processed on mitochondrial import (removing the first 58 residues) to generate the mature form [38]. Recent years have seen the determination of the atomic structures of bovine AdR and Adx [39–41]. The AdR structure reveals a significant charge separation on the surface, with the cleft between the FAD and NADP(H) binding segments of the protein being highly basic, whereas the opposite face of the protein is mainly acidic. This is considered to facilitate electrostatic interactions with the Ad protein, which is negatively charged in the region around its 2Fe-2S cluster [39, 41]. This is consistent with mutagenesis studies and effects of ionic strength on the interactions between these partners [42, 43]. Subsequent atomic structural analysis of a cross-linked complex between the bovine proteins revealed that Adx binds to the expected region of AdR and that some reorientation of the FAD- and NADP-binding domains of AdR occurs to facilitate the docking. As was also expected, electrostatic interactions appear to be a predominant force driving the binding of these partners. Burial of the Adx 2Fe-2S cofactor and of key residues implicated in interactions with P450 partners occurs in the binary complex of AdR/Adx. This renders unlikely the possibility that electron transfer to P450s can occur in a tertiary complex between AdR/Adx/ P450, and supports the notion that Adx shuttles between its electron donor and acceptor partners [44]. 5.2.5 Cytochrome P450 Reductase

The class II P450 redox system exploits the FAD- and FMN-containing CPR enzyme, which favors NADPH as its coenzyme. Mammalian CPRs are membraneassociated enzymes, with N-terminal anchor regions retaining these proteins in cellular membranes, and with the catalytic domains facing intracellularly [45]. All membranous hepatic P450s are supported by CPR. Analysis of the amino acid sequence of CPR enzymes indicates that they have two major domains (FAD/ NADPH-binding and FMN-binding) that have evolved from the fusion of genes encoding NADP(H)-ferredoxin reductase-like and flavodoxin-like proteins [46]. Removal of the N-terminal anchor region of the protein (either by proteolysis or protein engineering) enables production of a soluble form of the microsomal CPR [47]. The atomic structure for solubilized rat CPR confirms this domain organization and also demonstrates that the FAD and FMN flavins are closely juxtaposed to enable efficient inter-cofactor electron transfer [48]. As with the P450BM-3 system (see Section 5.2.6), the component FAD/NADPH and FMN domains have been expressed and purified, and shown to have redox and other properties consistent with those in the intact enzyme [49]. In CPR, NADPH reduces the FAD cofactor by hydride (2-electron) transfer, and electrons are transferred to the P450 via the FMN cofactor. Thus, the FMN acts as a single electron shuttle between FAD and heme. Potentiometric studies are consistent with this model, with the FMN being the more positive potential flavin

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[50]. Also, the relative potentials of the FMN oxidized/semiquinone and semiquinone/hydroquinone couples are consistent with the hydroquinone FMN as the electron donor to the heme iron, and with a model in which the enzyme undergoes a 1-3-2-1 cycle, where the digits refer to the number of electrons on the CPR flavins [51]. CPR is considered to be in a single electron reduced state at the start of the cycle, with the FMN in a semiquinone state. Reduction by NADPH places a further two electrons on the enzyme, with the FMN then being reduced to hydroquinone and the FAD to its semiquinone state on electronic equilibration. Both flavosemiquinones (FAD and FMN) are the blue (neutral) form. The first FMN-to-heme electron transfer from FMN hydroquinone is followed by redistribution of electrons in the CPR to reform the FMN hydroquinone and oxidize the FAD to its quinone state. The second FMN-to-heme electron transfer restores the starting (FMN semiquinone) state [52]. Eukaryotic CPR enzymes are often purified in a single electron reduced (FMN semiquinone) form, referred to as the “air stable semiquinone” state [53]. Protein engineering studies on CPR (predominantly rabbit, human, and rat isoforms) have demonstrated important roles for a “catalytic triad” of residues (Cys629, Asp674, and Ser457 in the human enzyme) in CPR that are crucial to efficient binding and electron transfer from NADPH [54]. Other CPR-like proteins include methionine synthase reductase, the reductase domain of nitric oxide synthase and the human cancer-related enzyme novel reductase 1 (NR1) [55–57]. NR1 has the slowest rate of flavin (FAD) reduction of all these enzymes and this is the rate-limiting step in catalysis for the enzyme [58]. In NR1, two of the three catalytic triad residues are mutated to other amino acids (Ala549 corresponds to Cys629, and Glu594 to Asp674 in human CPR), and this is almost certainly the reason for its slow kinetics. Other important mutations made to human CPR were the removal of the aromatic “lid” over the FAD cofactor in W676A/H variants. These effected substantial switches in coenzyme selectivity from NADPH to NADH, as discussed in Section 5.5 [59]. Several studies (modeling, chemical modification, and mutagenesis) of the interactions between microsomal CPR and its P450 partners point to the importance of acidic (Glu and Asp) residues in the FMN domain and basic residues on the P450 partners in electrostatic binding interactions [51, 60–64]. However, these data contrast with others demonstrating increased rates of electron transfer between CPR and P450s as ionic strength is increased, leading to disfavoring of ionic interactions [51, 65–67]. Plausible explanations include the possibility than alterations in the nature of interdomain (FAD/NADPH and FMN) interactions in CPR occur at elevated ionic strength so as to promote electron transfer in the systems, or that hydrophobic interactions that “fi ne tune” configurations that enhance interprotein electron transfer take place under such conditions. Much work clearly remains to be done to understand fully the nature of CPR interactions with its multiple P450 partners in cell membranes. In the 1980s it became clear that CPR was not an exclusively eukaryotic enzyme, and the characterization of microbial P450–CPR fusion enzymes provided


an important leap forward in the characterization of the P450 superfamily, as discussed in the section below. 5.2.6 P450BM-3 and Related CPR Fusion Enzymes

Armand Fulco’s studies of fatty acid hydroxylase activity in the soil bacterium Bacillus megaterium revealed the first major P450 redox system deviating from the aforementioned class I and class II types [14]. The hydroxylase activity was found to be associated with a phenobarbital-inducible 119 kDa protein, which was then shown to be a P450 (N-terminal) fused to a CPR module (C-terminal) [68, 69]. The enzyme (P450BM-3 or CYP102A1) is a soluble enzyme and a membrane anchor is absent from both the P450 and reductase domains of the enzyme. Thus, P450BM-3 was the first P450 shown to use a soluble CPR enzyme and the first natural P450/redox partner fusion enzyme [14, 70] (Fig. 5.3a). P450BM-3 has the highest catalytic center activity reported for a P450 monooxygenase system (>15 000 min−1 with arachidonic acid), and a highly efficient intra-reductase domain and reductase-to-heme electron transfer system underlies this rapid fatty acid hydroxylase rate [71, 72]. P450BM-3 catalyzes hydroxylation of a wide range of fatty acids (both saturated and unsaturated), typically hydroxylating close to the ϑ-terminal at the ω-1 to ω-3 positions [73]. As with P450cam, extensive protein engineering studies have revealed various aspects of structure and mechanism in the enzyme. Mutagenesis studies have revealed key roles for the active site residues Phe87 (control of regioselectivity of substrate oxygenation), Arg47 and Tyr51 (interactions with substrate carboxylate group), and Thr268 (oxygen activation and coupling of electron transfer to substrate oxidation) [71, 74, 75]. Key roles for Ala264 (mutants perturb protein conformational equilibria) and Phe393 (interactions with Cys–Fe bond and control of thermodynamic properties of the P450 heme iron) have also been established [76, 77]. In the reductase domain, the “catalytic triad” of residues (conserved across the diflavin reductase family of enzymes) in P450BM-3 comprises residues Cys999, Ser830, and Asp1044. A C999A mutant of P450BM-3 reductase was shown to be dramatically kinetically compromised as a result of a much slower rate of hydride transfer from the NADPH coenzyme and through formation of a stable NADP+ (i.e. product) complex [78]. However, other mutations conducive to the biotechnological application of P450BM-3 have also been reported, and are discussed in more detail in Section 5.4. Genetic dissection of the P450BM-3 flavocytochrome enabled production of catalytically viable P450 and reductase domains, and led to the production of FAD/NADPH- and FMN-binding domains of the reductase. These experiments validated the proposed domain composition of the flavocytochrome, and were important in confirming that the CPR module resulted from an ancestral fusion of genes encoding ferredoxin reductase-like and flavodoxin-like proteins [79, 80]. Domain dissection studies also enabled determination of reduction potentials for all of the redox centers in the enzyme, and facilitated the demonstration that fatty

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Fig. 5.3 Novel types of P450 systems. Several types of P450 systems distinct from those class I and class II systems represented in Fig. 5.1 are now known. (a) P450BM-3 (CYP102A1) type system, with P450 fused to a CPR. The structures shown are the BM-3 heme domain (PDB code 2HPD) and rat CPR (1AMO). (b) CYP116B1/B2 type system with P450 fused to a PDOR enzyme. Representative structures shown are Saccharopolyspora erythrae P450 eryF (CYP107A1, PDB code 1OXA) and the phthalate dioxygenase reductase from Burkholderia cepacia (2PIA). (c) Methylococcus capsulatus CYP51–ferredoxin fusion, represented by M. tuberculosis CYP51 (1X8V) and the Pyrococcus furiousus 3Fe-4S ferredoxin (1SJ1). (d) Rh. rhodochrous Y-11 XplA enzyme, considered to be a

P450–flavodoxin fusion. The structure is represented by rabbit CYP2B4 (2BDM) and E. coli flavodoxin (1AHN). (e) A P450 system that does not have a protein redox partner (representing the types of P450 driven by peroxide or by direct interaction with NAD(P)H). The protein shown is Fusarium oxysporum P450nor, with bound nitric oxide (1CL6). (f) Likely Ps. fluorescens P450-acyl CoA dehydrogenase fusion represented by Polyangium cellulosum P450 epoK (1Q5E) and Sus scrofa acyl CoA dehydrogenase (3MDE). (g) Non-NAD(P)H-dependent P450 system from Sulfolobus solfataricus, represented by the Desulfovibrio africanus pyruvate ferredoxin oxidoreductase (2C3M), the seven-iron ferredoxin from Azotobacter vinelandii (1FD2) and Sulfolobus solfataricus CYP119A1 (1F4U).

acid substrate binding leads to an increase in heme iron redox potential that triggers NADPH-dependent electron transfer from the redox partner [13]. Atomic structures of the P450 (heme) domain have been solved in substratefree and substrate-bound (both palmitoleic acid and N-palmitoylglycine) forms. These reveal considerable conformational alterations [81–83]. It remains unclear whether these major structural rearrangements are a consequence of substrate


binding per se, or instead reflect natural conformations of the ligand-free enzyme that may have differing affinities for substrates. Recent structural studies on an A264E mutant of the BM-3 heme domain suggest the latter explanation has some merit [84]. While there is, as yet, no atomic structure for the CPR domain of the enzyme, the FMN domain has been solved in a non-stoichiometric complex with the heme domain [85]. The FMN domain structure highlights important structural differences between the BM-3 FMN domain and other bacterial flavodoxins that might underlie its tendency to form an anionic (red) as opposed to a neutral (blue) semiquinone species [86, 87]. The BM-3 FMN–heme complex structure located the FMN domain near the proximal face of the heme (the likely docking interface) with its flavin cofactor orientated towards the heme cysteinate ligand [85]. However, a predicted electron transfer pathway through ∼50 sigma bonds can be ruled out as a consequence of the discrepancy between the measured and predicted rates of this process [14, 72]. Mobility of the FMN domain (between its electron donor FAD domain and acceptor heme domain) of the enzyme is likely to occur to bring cofactors into close proximity for efficient electron transfer. However, recent data indicate that the flavocytochrome is catalytically functional in fatty acid hydroxylation as a dimer, with electron transfer between FMN in one monomer and heme in the other supporting fatty acid hydroxylation [88]. Thus, a more detailed understanding of the interdomain interactions likely awaits the determination of the atomic structure of the intact flavocytochrome enzyme. The physiological role of P450BM-3 remains uncertain, but recent genome sequencing efforts have confirmed its presence in several other bacteria – including Ralstonia metallidurans, Bacillus subtilis, and Bradyrhizobium japonicum. In B. subtilis there are two homologs – CYP102A2 and CYP102A3 [16]. Both the A2 and A3 enzymes catalyze fatty acid hydroxylation close to the ϑ-terminal of fatty acid substrates, as does P450BM-3. However, the properties of these enzymes deviate from BM-3 and from one another in terms of substrate preference (K d values) and turnover rates (kcat values), and with respect to apparent cooperative binding (sigmoidal binding curves) observed for various lipid substrates. The A3 enzyme has been successfully engineered to produce 1-octanol from octane, and it is clear that the catalytic diversity of the CYP102 enzyme class can be exploited for biotechnologically important reactions [89]. Amino acid sequence alignment of B. megaterium P450BM-3 with its B. subtilis homologs reveals important regions of conservation and deviation between these enzymes (Fig. 5.4). Among the most notable features are the absences of residues corresponding to BM-3 Arg47 and Tyr51 in the A2/A3 enzymes. These residues are implicated in binding fatty acid carboxylate in the BM-3 enzyme, but are clearly not conserved for such a role in A2/A3 [71]. The strong conservation of the I helix region (indicated for the BM-3 enzyme in blue text in Fig. 5.4a) in the A2/A3 enzymes is consistent with the similar substrate selectivity and reactivity of these three enzymes (Fig. 5.4a). In the reductase domain, regions involved in binding coenzyme (NADPH) pyrophosphate and FAD cofactor are

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5 Cytochrome P450 Redox Partner Systems: Biodiversity and Biotechnological Implications CYP102A2 CYP102A3 CYP102A1

MKETSPIPQPKTFGPLGNLPLIDKDKPTLSLIKLAEEQGPIFQIHTPAGTTIVVSGHELV 60 MKQASAIPQPKTYGPLKNLPHLEKEQLSQSLWRIADELGPIFRFDFPGVSSVFVSGHNLV 60 --TIKEMPQPKTFGELKNLPLLNTDKPVQALMKIADELGEIFKFEAPGRVTRYLSSQRLI 58 . :*****:* * *** ::.:: :* ::*:* * **::. *. : :*.:.*:

CYP102A2 CYP102A3 CYP102A1

KEVCDEERFDKSIEGALEKVRAFSGDGLFTSWTHEPNWRKAHNILMPTFSQRAMKDYHEK 120 AEVCDEKRFDKNLGKGLQKVREFGGDGLFTSWTHEPNWQKAHRILLPSFSQKAMKGYHSM 120 KEACDESRFDKNLSQALKFVRDFAGDGLFTSWTHEKNWKKAHNILLPSFSQQAMKGYHAM 118 *.***.****.: .*: ** *.*********** **:***.**:*:***:***.**


MVDIAVQLIQKWARLNPNEAVDVPGDMTRLTLDTIGLCGFNYRFNSYYRETPHPFINSMV 180 MLDIATQLIQKWSRLNPNEEIDVADDMTRLTLDTIGLCGFNYRFNSFYRDSQHPFITSML 180 MVDIAVQLVQKWERLNADEHIEVPEDMTRLTLDTIGLCGFNYRFNSFYRDQPHPFITSMV 178 *:***.**:*** ***.:* ::*. *********************:**: ****.**:


RALDEAMHQMQRLDVQDKLMVRTKRQFRYDIQTMFSLVDSIIAERRANGDQDEKDLLARM 240 RALKEAMNQSKRLGLQDKMMVKTKLQFQKDIEVMNSLVDRMIAERKANPDENIKDLLSLM 240 RALDEAMNKLQRANPDDPAYDENKRQFQEDIKVMNDLVDKIIADRKASGEQS-DDLLTHM 237 ***.***:: :* . :* ..* **: **:.* .*** :**:*:*. ::. .***: *


LNVEDPETGEKLDDENIRFQIITFLIAGHETTSGLLSFATYFLLKHPDKLKKAYEEVDRV 300 LYAKDPVTGETLDDENIRYQIITFLIAGHETTSGLLSFAIYCLLTHPEKLKKAQEEADRV 300 LNGKDPETGEPLDDENIRYQIITFLIAGHETTSGLLSFALYFLVKNPHVLQKAAEEAARV 297 * :** *** *******:******************** * *:.:*. *:** **. **


LTDAAPTYKQVLELTYIRMILNESLRLWPTAPAFSLYPKEDTVIGGKFPITTNDRISVLI 360 LTDDTPEYKQIQQLKYIRMVLNETLRLYPTAPAFSLYAKEDTVLGGEYPISKGQPVTVLI 360 LVDPVPSYKQVKQLKYVGMVLNEALRLWPTAPAFSLYAKEDTVLGGEYPLEKGDELMVLI 357 *.* .* ***: :*.*: *:***:***:*********.*****:**::*: ..: : ***


PQLHRDRDAWGKDAEEFRPERFEHQDQVPHHAYKPFGNGQRACIGMQFALHEATLVLGMI 420 PKLHRDQNAWGPDAEDFRPERFEDPSSIPHHAYKPFGNGQRACIGMQFALQEATMVLGLV 420 PQLHRDKTIWGDDVEEFRPERFENPSAIPQHAFKPFGNGQRACIGQQFALHEATLVLGMM 417 *:****: ** *.*:*******. . :*:**:************ ****:***:***::


LKYFTLIDHENYELDIKQTLTLKPGDFHISVQSRHQEAIHADVQAAEKAAPDEQKEKTEA 480 LKHFELINHTGYELKIKEALTIKPDDFKITVKPRKTAAINVQRKEQADIKAETKPKETKP 480 LKHFDFEDHTNYELDIKETLTLKPEGFVVKAKSKKIPLGGIPSPSTEQSAKKVRKKAENA 477 **:* : :* .***.**::**:** .* :..:.:: . . : : :.

Fig. 5.4 Amino acid alignment of Bacillus megaterium P450 BM3 (CYP102A1) and Bacillus subtilis CYP102A2 and CYP102A3. Various key residues recognized in studies of the P450 BM3 enzyme are highlighted in white text on a black background. Panel A shows an alignment of the heme domains of the three enzymes. Arg 47 and Tyr 51 are considered important for interactions with fatty acid carboxylate groups, but are not retained in CYP102A2 or A3, suggesting a differing substrate interaction mode [71]. Phe 87 is a determinant of regioselectivity of fatty acid hydroxylation in P450 BM3, and is retained in A2/A3 [71, 74]. Mutation of Leu 181 to Arg/Lys led to improved binding of

short chain fatty acids in BM3, and this residue is retained in A2/A3 [109]. The I helix residues Ala 264 and Thr 268 are retained in all enzymes, and the entire I helical region (bold text coloured in black and underlined for the BM3 enzyme) is very strongly conserved in these enzymes. In BM3, an A264E mutant produced a novel Cys-Fe-Glu heme iron ligand set [76]. Thr 268 is implicated in oxygen binding/proton delivery [75]. Cys 400 is the thiolate ligand to the heme iron and is retained in all P450s. Phe 393 is retained in A2/A3 and in all P450 monooxygenase enzymes, and is a critical determinant of heme iron reduction potential and oxy complex stability [13, 77].

5.2 P450 Redox Partners CYP102A2 CYP102A3 CYP102A1

KGASVIGLNNRPLLVLYGSDTGTAEGVARELADTASLHGVRTKTAPLNDRIGKLPKEGAV 540 K-------HGTPLLVLFGSNLGTAEGIAGELAAQGRQMGFTAETAPLDDYIGKLPEEGAV 533 H--------NTPLLVLYGSNMGTAEGTARDLADIAMSKGFAPQVATLDSHAGNLPREGAV 529 : . *****:**: ***** * :** . *. .:.*.*:. *:**.****


VIVTSSYNGKPPSNAGQFVQWLQEIKPGELEGVHYAVFGCGDHNWASTYQYVPRFIDEQL 600 VIVTASYNGAPPDNAAGFVEWLKELEEGQLKGVSYAVFGCGNRSWASTYQRIPRLIDDMM 593 LIVTASYNGHPPDNAKQFVDWLDQASADEVKGVRYSVFGCGDKNWATTYQKVPAFIDETL 589 :***:**** **.** **:**.: . .:::** *:*****::.**:*** :* :**: :


AEKGATRFSARGEGDVSGDFEGQLDEWKKSMWADAIKAFGLELNENADKE-RSTLSLQFV 659 KAKGASRLTAIGEGDAADDFESHRESWENRFWKETMDAF--DINEIAQKEDRPSLSITFL 651 AAKGAENIADRGEADASDDFEGTYEEWREHMWSDVAAYFNLDIENSEDNK--STLSLQFV 647 *** .:: **.*.:.***. :.*.: :* :. * :::: ::: .:**: *:


RGLGESPLARSYEASHASIAENRELQSADSDRSTRHIEIALPPDVEYQEGDHLGVLPKNS 719 SEATETPVAKAYGAFEGIVLENRELQTAASTRSTRHIELEIPAGKTYKEGDHIGILPKNS 711 DSAADMPLAKMHGAFSTNVVASKELQQPGSARSTRHLEIELPKEASYQEGDHLGVIPRNY 707 : *:*: : * : .:*** . * *****:*: :* *:****:*::*:*


QTNVSRILHRFGLKGTDQVTLSASGRSAGHLPLGRPVSLHDLLSYSVEVQEAATRAQIRE 779 RELVQRVLSRFGLQSNHVIKVSGSAHMA-HLPMDRPIKVVDLLSSYVELQEPASRLQLRE 770 EGIVNRVTARFGLDASQQIRLEAEEEKLAHLPLAKTVSVEELLQY-VELQDPVTRTQLRA 766 . *.*: ****.... : :... . ***: :.:.: :**. **:*:..:* *:*


LASFTVCPPHRRELEELS-AEGVYQEQILKKRISMLDLLEKYEACDMPFERFLELLRPLK 838 LASYTVCPPHQKELEQLVSDDGIYKEQVLAKRLTMLDFLEDYPACEMPFERFLALLPSLK 830 MAAKTVCPPHKVELEALL-EKQAYKEQVLAKRLTMLELLEKYPACEMKFSEFIALLPSIR 825 :*: ******: *** * . *:**:* **::**::**.* **:* *..*: ** .::


PRYYSISSSPRVNPRQASITVGVVRGPAWSGRGEYRGVASNDLAERQAGDDVVMFIRTPE 898 PRYYSISSSPKVHANIVSMTVGVVKASAWSGRGEYRGVASNYLAELNTGDAAACFIRTPQ 890 PRYYSISSSPRVDEKQASITVSVVSGEAWSGYGEYKGIASNYLAELQEGDTITCFISTPQ 885 **********:*. . .*:**.** . **** ***:*:*** *** : ** . ** **:


SRFQLPKDPETPIIMVGPGTGVAPFRGFLQARDVLKREGKTLGEAHLYFGCRN-DRDFIY 957 SGFQMPNDPETPMIMVGPGTGIAPFRGFIQARSVLKKEGSTLGEALLYFGCRRPDHDDLY 950 SEFTLPKDPETPLIMVGPGTGVAPFRGFVQARKQLKEQGQSLGEAHLYFGCRSPHEDYLY 945 * * :*:*****:********:******:***. **.:*.:**** ****** ..* :*


RDELERFEKDGIVTVHTAFSRKEGMPKTYVQHLMADQADTLISILDRGGRLYVCGDGSKM 1017 REELDQAEQDGLVTIRRCYSRVENEPKGYVQHLLKQDTQKLMTLIEKGAHIYVCGDGSQM 1010 QEELENAQSEGIITLHTAFSRMPNQPKTYVQHVMEQDGKKLIELLDQGAHFYICGDGSQM 1005 ::**:. :.:*::*:: .:** . ** ****:: :: ..*: ::::*.::*:*****:*


APDVEAALQKAYQAVHGTGEQEAQNWLRHLQDTGMYAKDVWAGI 1061 APDVERTLRLAYEAEKAASQEESAVWLQKLQDQRRYVKDVWTGM 1054 APAVEATLMKSYADVHQVSEADARLWLQQLEEKGRYAKDVWAG- 1048 ** ** :* :* : ..: :: **::*:: *.****:*

Fig. 5.4 Continued Panel B shows an alignment of the reductase domains of the three P450 systems. The gap in the A1/A3 alignments at the start indicates a different inter-domain linker region in the A2 enzyme. Residues Gly 570 and Trp 574 are important determinants of FMN binding and are retained in each enzyme [142]. Residues Cys 999, Ser 830 and Asp 1044 comprise a catalytic triad important for efficient electron transfer from NADPH coenzyme. They are retained in each enzyme and across the majority of the diflavin reductases [54, 78]. Ser 830 and Tyr 829 are located in a strongly conserved region involved in binding the FAD cofactor (bold text coloured in black and underlined for the BM3 enzyme), and the

side chain of Tyr 829 interacts with the FAD isoalloxazine ring system. The region between amino acids ~890–915 in BM3 encompasses a conserved region involved in interactions with NADP(H) and its pyrophosphate group, and is also coloured black in bold, underlined text) for the BM3 enzyme. Trp 1064 is also conserved in each enzyme, and the side chain of Trp 1064 stacks across the isoalloxazine ring system of the BM3 FAD cofactor and must be displaced to enable hydride ion transfer from NADPH. The W1064A/H mutants of P450 BM3 demonstrate a spectacular switch of coenzyme specificity in favour of NADH [127].

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also strongly conserved in A1–A3, as are the catalytic triad of residues essential for electron transfer from NADPH: Ser830, Cys999, and Asp1044 in BM-3 [54] (Fig. 5.4b). The CYP102-type fatty acid hydroxylase–CPR type of fusion protein is not restricted to bacteria, and P450foxy (CYP505A1 from the fungus Fusarium oxysporum) was the first eukaryotic representative of this class of enzymes characterized [90]. This protein associates with the cell membrane, but is devoid of any welldefined membrane anchor region. CYP505A1 also hydroxylates fatty acids at the ω-1, ω-2, and ω-3 positions. Similar enzymes are identifiable from sequence analysis of other eukaryotic genomes (e.g. Neurospora crassa), and CYP505B1 from Fusarium verticilloides likely functions as a polyketide hydroxylase in synthesis of the mycotoxin fumonisin [91]. 5.2.7 A Novel Class of P450–Redox Partner Fusion Enzymes

While the BM-3-type P450–CPR fusion system represented a novel class of P450 protein, the use of CPR enzymes to drive P450 catalysis was already well established. However, genome sequence data again alerted the community to a distinctive type of P450–redox partner fusion enzyme thus far found in Rhodococcus erythropolis, a number of species of the pathogen Burkholderia, and in R. metallidurans. This enzyme has a P450 (N-terminal) fused to a C-terminal reductase domain resembling the well-characterized phthalate dioxygenase reductase (PDOR) enzyme that binds both FMN and 2Fe-2S cofactors and which transfers electrons to phthalate dioxygenase (PDO) [92] (Fig. 5.3b). The PDO from Burkholderia cepacia catalyzes the first step in the breakdown of the aromatic compound phthalate, forming a cis-dihydrodiol product [93]. Subsequent conversion by pathway enzymes leads to protocatechuate (3,4-dihydroxybenzoate), an intermediate in aromatic metabolic pathways [94]. The PDOR is reduced preferentially by NADH (over NADPH) and electrons are transferred through the FMN, then 2Fe-2S centers and onto the PDO [95]. The first such P450–PDOR fusion enzyme characterized originated from Rhodococcus sp. NCIMB 9784 [96]. The enzyme (CYP116B2) has been expressed in E. coli and purified, and its individual domains have also been isolated [96, 97]. While a bona fide physiological substrate has not been defined for this enzyme, it was shown to dealkylate 7-ethoxycoumarin at a low rate. This is an assay used widely for demonstrating P450 monooxygenase activity. The heme (P450) domain of this fusion P450 has >50 % amino acid identity with the class I P450 ThcB (CYP116A1) from Rhodococcus erythropolis. The gene encoding this class I P450 is adjacent to genes that likely encode its physiological redox partners – a ferredoxin reductase (rhodocoxin reductase, thcD) and a 2Fe-2S ferredoxin (rhodocoxin, thcC). CYP116A1 catalyzes the oxidative degradation of atrazine and thiocarbamate-based herbicides, including EPTC (S-ethyl dipropylthiocarbamate) [98]. CYP116B2 does not appear to have herbicide oxidation activity, but our recent studies of another member of the P450–PDOR fusion class, CYP116B1 from R.

5.3 Increasing P450–Redox Partner Complexity: Flavodoxins and Diverse Ferredoxins

metallidurans, indicate that this enzyme does catalyze such reactions (Warman, A.J. et al., unpublished data). Research on this novel type of P450–redox partner fusion class is in its infancy, but genome analysis indicates that such enzymes are also present in various species of the pathogenic bacterium Burkholderia, and thus may be as similarly widespread in nature as are the CYP102 class of P450–CPR fusions. The biotechnological potential of such enzymes will not be clear until more detailed analysis of their substrate selectivity, stability, and efficiency of their electron transport systems is undertaken. However, they may function in resistance to environmental toxins, and thus could be exploited in, for example, plant engineering for herbicide resistance.

5.3 Increasing P450–Redox Partner Complexity: Flavodoxins and Diverse Ferredoxins

The requirement for a single electron donor system for the P450s is met by ferredoxin enzymes of the 2Fe-2S class in the mitochondrial adrenodoxin system, and in the well-characterized bacterial P450cam/putidaredoxin system. However, ferredoxins with other types of iron–sulfur clusters are found in nature, and the 3Fe-4S and 4Fe-4S ferredoxins are widespread in different organisms. Ferredoxins with these clusters have been shown to support P450 systems. For instance, the 4Fe-4S ferredoxin Fer was shown to drive fatty acid oxygenation catalyzed by Bacillus subtilis P450 BioI (CYP107H1) [99, 100]. Also, 3Fe-4S ferredoxins are known to support catalysis in, for example, the Mycobacterium tuberculosis CYP51 (sterol demethylase) P450 [101, 102]. In the M. tuberculosis system the gene encoding the 3Fe-4S ferredoxin (Fdx) is next to the P450 on the genome (genes Rv0763c and Rv0764c, respectively) and is cotranscribed. Ferredoxins and flavodoxins have similar (and frequently overlapping) functions in prokaryotes, for example in nitrogen fi xation reactions (e.g. 103). Flavodoxins have capacity to accept either one or two electrons, forming semiquinone (SQ) or hydroquinone (HQ) states, respectively. However, they usually act as single electron carriers in biological systems, often shuttling between HQ/SQ forms [104]. In many cases, the redox potentials of the flavodoxins’ HQ/SQ couples are also compatible with reduction of P450 heme [105]. Thus, it was not unexpected that flavodoxins might support catalysis of certain bacterial P450s, particularly in view of the evolutionary and structural relationships between flavodoxins and the FMN-binding (P450 electron donor) domains of eukaryotic and prokaryotic CPR modules [46]. In the B. subtilis BioI system, both host flavodoxins (YkuN and YkuP) support fatty acid oxygenation by the P450 (CYP107H1) [15, 106]. The flavodoxin cindoxin was also demonstrated to be the natural redox partner for P450 cin in Citrobacter braakii, and its gene is located chromosomally adjacent to that for its partner enzyme. P450 cin catalyzes cineole oxidation in a pathway enabling Citrobacter braakii to use cineole as an energy source for growth [107]. Pivotal studies in the Waterman laboratory also showed that mammalian

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P450s expressed in E. coli had their oxygenase activities supported by a system comprising the bacterium’s NADPH-flavodoxin reductase and flavodoxin proteins [e.g. 108]. The adaptability and evolvability of the prokaryotic P450s is manifest in their exploitation of a diverse range of cellular electron donor proteins. Key factors are clearly the evolution of a productive interaction site involving the proximal face of the P450, and the ability of the relevant flavodoxin/ferredoxin to donate single electrons to the heme iron at appropriate potentials.

5.4 Natural and Artificial P450–Redox Partner Fusion Enzymes and their Biocatalytic Potential

The biotechnological advantages offered by the fusion of P450s to redox partner enzymes appear clear from studies on the P450BM-3 system. BM-3 has an efficient electron transport chain that enables fatty acid hydroxylation at rates ∼100-fold faster than many eukaryotic P450s. In addition, it is catalytically self-sufficient, obviating the requirement for isolation of 2–3 different enzymes and determining ratios of redox partner-to-P450 that give optimal turnover [14, 72]. The same advantages are inherent in the A2/A3 systems and the other members of this class. While work is in its infancy for the CYP116B1/B2 systems, these are also catalytically self-sufficient and are likely to offer similar advantages in terms of efficient electron transfer apparatus. There are already several examples of the engineering of the P450BM-3 enzyme to enable its production of industrially relevant molecules. These include oxygenation of short-chain alkanoic acids, highly branched chain fatty acids (leading to chiral precursors for polyketide synthesis), and (+)valencene (to produce the grapefruit flavor compound (+)-nootkatone) [109–111]. CYP102A3 has also been engineered to enable enhanced production of 1octanol from octane [89]. Virtually all efforts at engineering the BM-3 enzyme have been directed at altering the structure of the heme domain to effect changes in substrate selectivity and product formation. However, there are also some serious issues to be addressed with respect to the performance of the reductase domain, as discussed below in Section 5.6. In the last few years (and as a direct result of the information generated from genome sequencing programs) new types of P450–redox partner fusions have been recognized, and a number have been expressed and characterized at the protein level. In Methylococcus capsulatus a P450 (N-terminal)–ferredoxin fusion protein occurs, and the P450 is clearly a member of the sterol demethylase (CYP51) family. The ferredoxin is likely to bind a 3Fe-4S cluster [112] (Fig. 5.3c). It appears likely that this system has evolved to fuse the P450’s cognate ferredoxin, and now requires only a separate ferredoxin reductase to complete the novel form of a class I P450 redox system. An analogous system was recognized in the bacterium Rhodococcus rhodochrous (strain Y-11), in which a flavodoxin (N-terminal) is fused to a soluble P450 (Fig. 5.3d). The enzyme (XplA) was shown to degrade

5.4 Natural and Artificial P450–Redox Partner Fusion Enzymes and their Biocatalytic Potential

the explosive RDX (Royal Demolition eXplosive – the molecule hexahydro-1,3,5trinitro-1,3,5-triazine) – although it is currently unclear whether the process involves oxygenase activity of the P450, or simply substrate reduction [113]. As with the M. capsulatus enzyme, a reductase enzyme is clearly required to complete the Rh. rhodochrous P450–redox system, and work on this explosive degrading enzyme is clearly aimed at producing a plant-based system for degrading RDX in the soil (phytoremediation). While not strictly a cytochrome P450, the similarity of the multidomain eukaryotic nitric oxide synthase (NOS) enzymes to P450BM-3 (and its homologs) cannot be overlooked in the context of oxygenase fusion enzymes. The major mammalian NOS isoforms (neuronal, endothelial, and inducible) all have a thiolate-coordinated heme enzyme (N-terminal) fused to a CPR module [114]. The enzymes are functional as dimers (as P450BM-3 is in fatty acid hydroxylation), and electron transfer occurs between the reductase of monomer 1 and the heme domain of monomer 2 in the dimer [88, 115]. The similarities between NOS enzymes and the BM-3 class of P450s extend to the fact that NOS enzymes perform hydroxylation/oxygenation chemistry. They hydroxylate the substrate L-arginine to N-hydroxy-L-arginine, and then (in a second reaction on this product) go on to form L-citrulline and nitric oxide (NO) at the heme center. However, there are important differences between NOS and BM-3 (and other P450s). The protein effector calmodulin plays important regulatory roles in the control of electron transfer in NOS, and the cofactor tetrahydrobiopterin (H4B) is bound close to the heme and also participates in catalytic redox reactions [114]. In addition, while the reductase module is structurally closely related to rat CPR, the oxygenase domain has a considerably different structure to the P450s, with a much more solvent exposed active site (Fig. 5.5). Notwithstanding the important differences between the NOS and P450–CPR structures, attempts have been made to create heterologous fusion proteins involving the different domains. In studies of fusions between BM-3 and rat neuronal NOS, catalytically functional chimeras were generated – with the most successful fusion being that between the BM-3 reductase domain and the nNOS heme domain, which catalyzed NO production quite efficiently [116]. Numerous attempts have also been made to mimic the natural BM-3-type P450 fusion systems by fusing eukaryotic P450s with CPR. Frequently, some enhancement of activity is obtained – although no fusions created to date have produced chimeras with activity levels that approach those of P450BM-3 or its A2/A3 relatives. Examples of stable, active P450–CPR fusions reported include ones involving the mammalian P450s CYP3A4 and CYP17A [117, 118]. Other interesting fusion proteins generated included three-protein chimeras involving P450cam and its PdR and Pd redox partners. The most efficient chimera created (a PdR–Pd–P450 chimera from N- to C-terminus) had a kcat for camphor hydroxylation of ∼30 min−1, around 100-fold lower than can be obtained by reconstituting the isolated redox partner proteins [119]. Thus, artificial P450–redox partner fusion protein constructs have provided useful tools to enable production of single-component, catalytically self-sufficient

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Fig. 5.5. Atomic structures of the oxygenase and reductase domains of eukaryotic nitric oxide synthase. (a) Atomic structure of murine iNOS bound to L-arginine (PDB code 1DWV) [147]. The distinctly different protein fold to those of the P450s is obvious from comparison with the structures shown in Figs 5.1 and 5.3. The relatively exposed heme is shown at the center of the structure in red spacefill. L-Arginine is shown in colored ball and stick representation above the plane of the heme. (b) Atomic structure of rat neuronal NOS reductase. The overall structure shows strong similarities to that of rat CPR [48]. Domains of the NOS reductase protein are color coded as follows: NADPHbinding domain in red; FAD-binding domain

in dark blue; alpha helical connecting domain (CD, which orientates the two flavin cofactors) in cyan; and FMN-binding domain in orange. Other prominent features are the autoinhibitory helix (purple) at the bottom left of the structure that is located in the FMN-binding domain and which interferes with CaM binding and likely modulates both intra and inter-domain electron transfer; and the β-finger/CD2A section (green beta sheet section and associated yellow loop region) extending from the CD and which also influences regulation by Ca2+/CaM. The NADP+, FAD and FMN cofactors are shown in green, orange, and yellow stick representation [56].

protein entities, and have potential applications in biotechnology. However, their activity levels rarely exceed those achieved by reconstituting their separate component enzymes, and thus do little to explain the efficiency of the electron transfer systems seen in the “natural” (e.g. P450BM-3) P450–redox partner fusion enzymes.

5.5 Other Routes to Driving P450 Catalytic Function

The fact that most P450 enzymes require at least one redox partner protein is often seen as a complicating factor in their exploitation for biotechnological applications. However, there are means to drive P450 catalysis that do not rely on exogenous protein partners. The best known of these is the peroxide shunt mechanism (as illustrated in Fig. 5.2) whereby the resting state of the P450 can be converted to the ferric hydroperoxy form by reaction with hydrogen peroxide (or organic peroxides). This is the reversal of the uncoupling reaction that leads

5.5 Other Routes to Driving P450 Catalytic Function

to collapse of the ferric hydroperoxy species. Protonation of this species creates the reactive compound I, although the ferric hydroperoxy state itself may also participate in certain P450-dependent reactions [18, 120]. In theory, the peroxide shunt represents a cheap alternative to use of expensive nicotinamide coenzymes and multiple protein partners. However, the reality is that peroxides oxidatively degrade the heme macrocycle and attack amino acid side-chains. Thus, P450 inactivation appears an inevitable consequence of exploitation of the peroxide shunt method, and this has prevented its widespread application for chemical transformations. Recent mutagenesis studies have aimed to address the peroxide instability issue, and have generated P450BM-3 mutants with moderately increased activity (but not stability) with peroxide [121]. Perhaps inevitably, nature has found more appropriate structural solutions to the problem, and B. subtilis P450BSβ (CYP 152A1) is a peroxygenase P450 that has evolved to use hydrogen peroxide directly to facilitate hydroxylation of fatty acids [122]. Homologs of this P450 have been recognized in other organisms (e.g. Sphingomonas paucimobilis, CYP152B1), and analysis of its atomic structure will clearly provide important clues as to adaptations that enhance its stability/activity with peroxide [123]. Electrocatalysis of P450s (i.e. providing electrons directly from an electrode) provides a further means by which P450 turnover can be induced in the absence of protein partners and nicotinamide coenzymes. As with the peroxide shunt, there are several drawbacks relating to such phenomena as (a) denaturation/ unfolding of the P450 at an electrode surface, (b) problems associated with providing temporally distinct electrons (enabling binding of dioxygen to ferrous heme iron prior to further reduction), and (c) turnover of P450 in absence of substrate, leading to production of damaging radicals. Notwithstanding these problems, there are examples of successful exploitation of electrocatalysis. For example, Estabrook and co-workers demonstrated turnover of rat CYP4A1 in a system that used the mediator cobalt sepulchrate to transfer electrons between an electrode and the dissolved P450 and rat CPR, or a rat CYP4A1–CPR fusion protein. Rates of formation of the natural hydroxylated fatty acid product (12-hydroxydodecanoic acid) were comparable with those achieved using NADPH as the electron donor [124]. Direct electrochemistry of immobilized human CYP2E1 has been demonstrated, and recent studies also reported reduction of P450BM-3 immobilized on a graphite electrode [125, 126]. Given problems with artificial routes to driving P450 catalysis, the cost implications relating to use of a coenzyme driven system may not be an overriding factor mitigating against exploiting such systems for biotransformations. Indeed, when the rapid turnover rates and tight coupling of electron transfer to substrate oxygenation demonstrated by the P450BM-3 (and P450cam) systems are considered, the operation of a “traditional” P450 turnover system may have distinct advantages over the aforementioned electrocatalytic and peroxide shunt systems. The expense of the coenzyme NADPH can be substantially offset by inclusion of a NADPH-regenerating system (e.g. glucose 6-phosphate dehydrogenase) in which NADP+ is reduced back to NADPH at the expense of exhausting a much cheaper

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chemical (glucose 6-phosphate). A similar system can be used to regenerate NADH from NAD + (e.g. glucose dehydrogenase and glucose). Protein engineering has also been used to make substantial changes in coenzyme specificity from NADPH towards the much cheaper NADH for both mammalian CPR and the BM-3 enzyme, with replacement of a FAD-shielding aromatic residue in both enzymes (W676A in human CPR and W1046A in P450BM-3) effecting huge switches in catalytic efficiency (kcat/K m) towards NADH (∼1000-fold and ∼5700fold, respectively) [59, 127]. Another exciting development in the P450 field has been the identification of a P450 system which uses NAD(P)H directly without requirement for any redox partners. The Fusarium oxysporum P450nor (CYP55A1) binds NADH directly to a P450 and uses electrons from the coenzyme to reduce two molecules of NO to N2O [128] (Fig. 5.3e). Structural studies are providing important insights into mechanistic aspects of this reaction, although it is not immediately obvious whether such a reductive system could be evolved into one capable of supporting P450 oxygenase chemistry [129].

5.6 Uncoupling, Enzyme Stability and Coenzyme Issues

As illustrated in Fig. 5.2, the cytochromes P450 are notorious for their tendency to undergo abortive catalytic cycles with the production of superoxide, peroxide, or water according to the point in the cycle at which the relevant oxy complex collapses. Obviously, the intracellular formation of peroxide and superoxide is undesirable and these radicals have potential to damage the P450 protein or heme macrocycle (leading to enzyme inactivation), or to escape the active site to react with other cellular molecules [130]. There are a number of potential reasons underlying these phenomena in various P450s. Following the first reductive step in the cycle, the ferrous oxy complex can decay to form superoxide if the second electron is not delivered efficiently to enable the progression of the cycle and the formation of more powerful oxidant species. The ferrous oxy complex is relatively short-lived in the P450cam enzyme, but is stabilized by the presence of substrate [131]. However, the complex has a much shorter lifetime in, for instance, P450BM3 [77], and its instability is clearly also a major issue underlying the uncoupling of several mammalian isoforms [120]. Many bacterial isoforms (e.g. P450BM-3 and P450cam) appear to have an excellent mechanism of avoiding non-productive electron transfer from the redox partner in absence of substrate. This occurs by substrate-dependent desolvation of the heme iron and conversion of the heme iron to a high-spin species with a concomitant positive shift in heme iron redox potential [12, 13]. This switch triggers the electron transfer process to heme iron only in the presence of substrate. This type of regulation appears much rarer in mammalian P450s, and various isoforms are isolated in mixed-spin or substantially high-spin forms even in absence of substrate. Electron transfer to a substrate-free P450 will inevitably lead to eventual decay of the ferrous oxy complex unless substrate can bind in advance

5.7 Future Prospects

of either oxy complex decay or further reduction (which will generate even shorter lived oxy species) [120]. A further potential reason for the collapse of the ferrous oxy or later intermediates in the cycle is the inappropriate positioning of the substrate in the active site (i.e. substrate location too distant from the oxy complex to enable it to be attacked and oxygenated). In addition, if the substrate-like molecule presents a bond that cannot be productively attacked (i.e. is too strong), then abortive decay of the ferryl-oxo (or ferric hydroperoxy) form should occur [18]. Obviously, the collapse of the ferryl-oxo species (with water formation) is less deleterious than peroxide production via ferric hydroperoxy collapse; but both mechanisms result in nonproductive oxidation of NAD(P)H coenzyme in the cell. A common conception is that the mammalian drug-metabolizing P450s are designed to have broad substrate specificity, enabling oxygenation of a wide range of physiological and xenobiotic substrates. Such broad specificity may be achieved by large, flexible active sites (as is clearly seen in the case of the human CYP3A4 enzyme) and frequent uncoupling of electron transfer from substrate oxygenation may be a necessary evil associated with having a P450 system capable of dealing with a plethora of different organic molecules [132]. By contrast, many bacterial P450s have very well-coupled electron systems and far more restricted substrate selectivity (e.g. the P450cam system) [8]. The major bacterial P450s (particularly P450cam and P450BM-3) also have much higher turnover rates than eukaryotic counterparts, and it is thus no surprise that such systems have been most popular in rational engineering and forced evolution experiments to create catalytically efficient P450 systems with desirable substrate recognition properties.

5.7 Future Prospects

Recent years have seen a dramatic increase in our knowledge of the biodiversity of cytochrome P450 redox systems. Aside from those systems mentioned above, other recent examples demonstrating the flexibility of redox partner selection in microbial P450s include the presence of a P450–acyl CoA dehydrogenase fusion protein system in Pseudomonas fluorescens (Fig. 5.3f) and a P450 enzyme (CYP119A1) from the thermophilic bacterium Sulfolobus solfataricus whose activity is supported by a 7Fe ferredoxin (likely comprising both 3Fe-4S and 4Fe-4S clusters, in which the 3Fe-4S cluster is redox active in catalysis) and a 2-oxoacid (pyruvate)-dependent ferredoxin oxidoreductase (Fig. 5.3g) [133, 134]. The thermophilic system has the obvious advantage that costly nicotinamide coenzymes could be avoided through use of pyruvate as the electron donor [133]. In terms of the use of protein engineering to alter the catalytic properties of P450 systems, the most broad ranging and successful studies have been done with the P450cam and P450BM-3 systems (and BM-3’s B. subtilis relatives CYP102A2 and A3), producing variants capable of efficient hydroxylation of such molecules as alkanes, polyunsaturated fatty acids and drugs, and with potential applications in, for example, the generation of high value lipid mediators, ethanol

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and authentic human metabolites of drug molecules [e.g. 135, 136]. Despite notable breakthroughs, it is still the case that P450s are thought of as relatively fragile biocatalysts, vulnerable to inactivation through structural disruption and/ or cofactor dissociation. In particular, the P450s are known to undergo a conversion from an “active” P450 form (in which the heme Soret band is positioned close to 450 nm in the ferrous carbon monoxy complex) to an “inactive” P420 form (with the peak near 420 nm). However, recent biochemical and spectroscopic studies indicate that such transitions, at least for soluble P450s, likely reflect reversible protonation of the heme thiolate (to a thiol) and that substrate binding stabilizes the P450 form [102, 137, 138]. Further stabilization of heme binding may be achieved by covalent binding of the heme macrocycle to the protein, which occurs naturally in eukaryotic CYP4 enzymes by turnover-dependent linkage of a heme methyl group to a conserved acidic (glutamate) residue in the I helix [139, 140]. This was also shown to occur partially in a variant of P450cam in which a glutamate was engineered into the I helix at the appropriate position [141]. Other stability issues associated with extended turnover of P450 redox systems include the loss or degradation of non-covalently bound cofactors from the redox partners. In the BM-3-type (CYP102 class) P450–CPR fusion enzymes, for instance, FMN is relatively weakly bound by comparison with homologous flavodoxin enzymes [142, 143]. Tightening binding of FMN in these enzymes is an obvious target for protein engineering studies, given the increasing interest in exploitation of the CYP102 (A1–A3) systems for production of chiral oxygenated molecules, and the requirement for extended turnover to facilitate high product yield. Many of the most biotechnologically attractive P450 systems are inactivated quite rapidly at temperatures above ∼40 ºC due to cofactor dissociation or irreversible structural change/aggregation [e.g. 143]. This has been addressed in theoretical, rational, and forced evolution studies of peroxygenase and other P450 systems, and also with reference to the known structural properties of P450s from thermostable organisms [e.g. 144, 145]. Clearly such studies may enable creation of more robust P450 catalysts that have greater useful lifetimes and higher thermostability. A similar approach was used to produce modest increases in catalytic efficiency of the P450BM-3 system in the peroxide shunt reaction [121]. However, it appears highly unlikely that this reaction could ever compete with the efficiency of the “natural” NADPH-driven reaction, particularly given the fact that NADPH recycling systems can substantially diminish operating costs in this system and since peroxide-mediated heme destruction leads to oxidative degradation of the enzyme. If progress in the next 50 years of P450 research is as great as in the first 50 years, then several further novel biocatalysts (likely catalytically efficient P450/ redox partner fusion enzymes) will be discovered and characterized, and protein engineering strategies to evolve activities and stabilize existing and novel enzymes will make these systems fully cost effective. This will allow the widespread exploitation of P450 oxygenases in biotechnological processes as efficient and safer alternatives to traditional organic synthesis methods.

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orientation. Biochemistry 2005, 44, 145 Maves, S.A., Sligar, S.G. Understanding 13914–13920. thermostability in cytochrome P450 by Limburg, J., LeBrun, L.A., Ortiz de combinatorial mutagenesis. Protein Sci Montellano, P.R. The P450cam G248E 2001, 10, 161–168. mutant covalently binds its prosthetic 146 Wester, M.R., Johnson, E.F., heme group. Biochemistry 2005, 44, Marques-Soares, C., Dijols, S., 4091–4099. Dansette, P.M., Mansuy, D., Stout, Klein, M.L., Fulco, A.J. Critical C.D. Structure of mammalian residues involved in FMN binding and cytochrome P450 2C5 complexed with catalytic activity in cytochrome diclofenac at 2.1 Å resolution: evidence P450BM-3. J Biol Chem 1993, 268, for an induced fit model of substrate 7553–7561. binding. Biochemistry 2003, 42, Munro, A.W., Lindsay, J.G., Coggins, 9335–9345. J.R., Kelly, S.M., Price, N.C. Analysis of 147 Ghosh, D.K., Crane, B.R., Ghosh, S., the structural stability of the Wolan, D., Gachhui, R., Crooks, C., multidomain enzyme flavocytochrome Presta, A., Tainer, J.A., Getzoff, E.D., P-450 BM3. Biochim Biophys Acta 1996, Stuehr, D.J. Inducible nitric oxide 1296, 127–137. synthase: role of the N-terminal betaSalazar, O., Cirino, P.C., Arnold, F.H. hairpin hook and pterin-binding Thermostabilization of a cytochrome segment in dimerization and P450 peroxygenase. ChemBiochem tetrahydrobiopterin interaction. EMBO J 2003, 4, 891–893. 1999, 18, 6260–6270.

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6 Steroid Hydroxylation: Microbial Steroid Biotransformations Using Cytochrome P450 Enzymes Matthias Bureik and Rita Bernhardt

6.1 Introduction

Hormonal steroids are a class of steroids which are derivatives of androstane, a C19 steroid which has the skeleton structure of gonane (four rings in all-trans configuration) with two additional methyl groups at positions 10β and 13β, respectively (Fig. 6.1). Their biological activity is determined by different hydroxyand keto-groups, by double bonds at varying positions and also by the presence or absence of a side-chain at the 17β-position. Other classes of steroids, such as bile acids and cardioactive steroid lactones, differ from hormonal steroids by the arrangement of the rings. From these basic structures numerous synthetic analogs are derived with a changed or improved biological activity profi le due to the introduction of additional double bonds or substituents. Steroid hormones regulate essential biological functions in humans and animals, since they act as mineralocorticoids, as glucocorticoids, or as sex hormones. Moreover, many steroid hormone derivatives display anabolic, antihormonal, antiinflammatory, antirheumatic, contraceptive, or sedative activities. Their use as drugs has therefore increased tremendously since 1949 (when Merck first introduced cortisone), and for many decades the overwhelming demand for steroids by the pharmaceutical industry has far exceeded the availability of these compounds from natural sources. Today, steroids represent one of the largest sectors in the pharmaceutical industry with worldwide markets in the region of US$10 billion. Although the total chemical synthesis of steroids by Woodward and colleagues [1] constituted a brilliant scientific achievement, it is not economically competitive, so for many decades most steroids have been industrially produced by hemisynthesis that mainly starts from β-sitosterol (or diosgenin and other phytosterols) and involves a varying number of sophisticated chemical and microbial bioconversion steps. Of course, microbial steroid biotransformations must compete with alternative chemical reactions on a cost basis, and with many chemical reactions

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Fig. 6.1 Basic structures of steroids. (a) Androstane with nomenclature of carbon atoms and rings. (b) Stereochemical arrangement of the four rings in the all-trans form and relative position of 11α- and 11β-substituents.

being economically feasible, this issue has considerably limited the number of steroid biotransformations that have been actually applied on an industrial scale. Indeed, microbial steps are often circumvented by using more complex chemical reactions provided they can be justified economically. Despite their inherent advantages, microbial reactions also display some specific shortcomings, including the formation of side products, yield variations caused by biological variability due to different batches of cells, and the low solubility of steroids in aqueous solutions. Recently, the total biosynthesis of steroids from simple carbon sources has been accomplished with genetically engineered yeasts [2], an achievement that might well constitute a paradigm shift as it opens up the road to chemistry-free steroid production. However, in this chapter we will not focus on pathway engineering but rather concentrate on the different types of single biotransformation reactions that have been performed so far. As the advantages and shortcomings of each biotransformation reaction are discussed, the feasibility of their combination should be borne in mind.

6.2 Cytochrome P450-Dependent Steroid Hydroxylase Systems

Cytochromes P450 got their name from both their hemoprotein character and from their unusual spectral properties, displaying a typical absorption maximum of the reduced CO-bound complex at 450 nm: cytochrome stands for a hemopro-

6.2 Cytochrome P450-Dependent Steroid Hydroxylase Systems

tein, P for pigment and 450 reflects the absorption peak of the CO complex at 450 nm. The ability of reduced P450 to produce an absorption peak at 450 nm upon CO binding is still used for the estimation of the P450 content [3]. These proteins are encoded by a superfamily of genes, convert a broad variety of substrates and catalyze a variety of interesting chemical reactions. This enzyme family is involved in the biotransformation of drugs, the bioconversion of xenobiotics, the bioactivation of chemical carcinogens, the biosynthesis of physiologically important compounds such as steroids, fatty acids, eicosanoids, fat-soluble vitamins, bile acids, the conversion of alkanes, terpenes, and aromatic compounds as well as the degradation of herbicides and insecticides. There is also a wide range of reactions catalyzed by cytochromes P450, such as carbon hydroxylation, hetero-atom oxygenation, dealkylation, epoxidation, aromatic hydroxylation, reduction, dehalogenation (reviewed in [4–6]). To date, more than 6000 different P450 genes have been cloned up to date (for details see: http://drnelson.utmem.edu/CytochromeP450.html). Members of the same gene family are defined as usually having ≥40% sequence identity with a P450 protein from any other family. Mammalian sequences within the same subfamily are always >55% identical. The numbers of individual P450 enzymes in different species differ significantly, the highest numbers observed so far being displayed in plants. Cytochromes P450 are external monooxygenases (mixed function oxidases) since they catalyze the incorporation of a single atom of molecular oxygen into a substrate with the concomitant reduction of the other atom to water. The monooxygenases are divided into two classes, the internal and the external monooxygenases. Internal monooxygenases extract two reducing equivalents from the substrate to reduce one atom of dioxygen to water, whereas external monooxygenases utilize an external reductant (see [7]). Cytochrome P450 systems catalyze the following reaction: R-H + O2 + NAD(P)H + H + → R-OH + H2O + NAD(P) + Details of the structure and chemistry of cytochromes P450 have recently been summarized in an excellent review [8] and for this reason will not be discussed here. It should be mentioned, however, that cytochromes P450 do not only catalyze monooxygenations, but also oxidase and peroxidase reactions. Variations of this scheme for the reaction mechanism of P450, however, occur with different P450 systems such as thromboxane and prostacyclin synthase, nitric oxide reductase (CYP55A1), and others. These different reaction types are, in addition to the main reaction cycle, the basis for the versatility of cytochromes P450. As mentioned above, cytochromes P450 are external monooxygenases, which implies that they need an external electron donor to transfer the electrons necessary for oxygen activation and the subsequent substrate hydroxylation. Two main classes of cytochromes P450 can be defined with respect to their electron supporting system, although other subclasses also exist [9]: the mitochondrial/bacterial type and the microsomal type (Fig. 6.2). Microsomal cytochromes P450 are

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Fig. 6.2 Schematic organization of different cytochrome P450 systems. (a) bacterial system, (b) mitochondrial system. (c) microsomal system, (d) self-sufficient CYP102 (P450BM-3). Figure slightly modified from [21].

membrane bound and accept electrons from a microsomal NADPH-cytochrome P450 reductase, containing FAD and FMN. All drug and xenobiotica metabolizing cytochromes P450 isolated so far and, in addition, CYP102 (P450BM-3) isolated from Bacillus megaterium, have been shown to belong to this class. CYP102 consists of a polypeptide chain with two different domains, one comprising the hemoprotein and the other containing an FAD- and FMN-dependent reductase. Most of the other bacterial cytochromes P450 belong to the second class. They are soluble and obtain the electrons necessary for the reaction mechanism from an NADH-dependent FAD-containing reductase via an iron–sulfur protein of the [2Fe-2S] type. Mitochondrial cytochromes P450 involved in the side-chain cleavage of cholesterol, the 11β-hydroxylation of 11-deoxycortisol, the production of aldosterone, and vitamin D biosynthesis also belong to the latter class. These cytochromes P450 are localized in the inner mitochondrial membrane, whereas

6.3 Native Microorganisms in Steroid Biotransformation

the [2Fe-2S] protein called adrenodoxin (Adx) in adrenal steroid hydroxylase systems is a soluble protein of the matrix. The FAD-containing reductase adrenodoxin reductase (AdR) is associated with the inner mitochondrial membrane. Interaction of the cytochromes P450 with their corresponding electron donors is a necessary prerequisite of the catalytic cycle. Its specificity guarantees a sufficient reaction rate of catalysis and likewise a discrimination between different potential donors and acceptors of electrons to protect the system from shunt reactions. Since in liver microsomes many different isoenzymes have to interact with only one type of reductase it has to be expected that the binding site for reductase is very similar or identical on various cytochromes P450. Salt bridges are responsible for the recognition of the reductase by the P450 and the correct orientation of both proteins to each other (for a review, see [10]). In addition to microsomal reductase, some microsomal cytochromes P450 are able to accept the second electron from cytochrome b5. Cytochrome b5 has also been shown to exert a differential stimulatory action, dependent upon both the form of cytochrome P450 and the reaction substrate [11, 12] (reviewed in [13, 14]). In mitochondrial steroid hydroxylases and in the camphor hydroxylating bacterial P450 (CYP101) system a charge-pair interaction mechanism has been demonstrated by chemical modification, site-directed mutagenesis studies, and structural data of electron transfer complexes (for review, see [15]). In addition, the Cterminal peptide of adrenodoxin [16], the residue Tyr82 [17], and the loop covering the iron–sulfur cluster [18] were shown to be of pivotal importance for redox partner interaction. Like microsomal reductase, the mitochondrial ferredoxin has also to deliver electrons to different cytochromes P450. From the available data, a shuttle model is favored, where the oxidized ferredoxin interacts first with the ferredoxin reductase to undergo reduction, with the formation of a Fe3+ /Fe2+ iron–sulfur cluster [15]. It dissociates from the reductase and then interacts with the respective cytochrome P450, where it delivers this electron before returning to the reductase and transfers the second electron to the P450. The mechanism of electron transfer between the components of the different cytochrome P450 systems, one of the fundamental problems in life sciences, is not yet well understood. Although it could be conclusively shown that posttranslational modifications can regulate these electron transfer reactions at least in some cases [19], we are only at the very beginning of this field of research as well. Since electron transfer to the P450 in some cases seems to be low and rate-limiting in P450 catalysis [20], engineering of this step could potentially, however, lead to significantly improved biocatalysts. The current status of the biotechnological use as well as the possibilities of genetic engineering of cytochromes P450 were recently reviewed [21].

6.3 Native Microorganisms in Steroid Biotransformation

Since its first description at the end of the nineteenth century [22], microbial steroid synthesis has been performed using a large variety of native microorgan-

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isms such as eubacteria, actinomycetes, fungi imperfecti, and more. In 1937, the development of technically useful microbial processes in steroid production began at Schering with the synthesis of testosterone. After the pharmacological effects of cortisol and progesterone became known, research in this field developed rapidly in the 1950s and led to the identification of a panel of different microorganisms that catalyze steroid hydroxylations, hydrogenations, dehydrogenations, and the splitting of carbon–carbon bonds (reviewed in detail by [23]). However, only a limited number of microbial transformations reported in the scientific literature have actually been applied technically. These comprise side-chain degradation reactions, Δ1,2-, Δ1,4-, 17β-dehydrogenation, and several hydroxylation reactions. Among the latter ones 11α-hydroxylation, 11β-hydroxylation, and 16α-hydroxylation, are of the highest commercial significance and are therefore discussed in detail below. Other steroid hydroxylation sites, including those at the positions 7α, 9α, 14α, and 15α, seem also to have potential for industrial exploitation. In addition, the 6β-hydroxylation of compound ML-236BNa to form the blockbuster drug pravastatin by Streptomyces carbophilus as well as the 1α- and 25-hydroxylations of the seco-steroid vitamin D3 by Amycolata species are prominent examples of P450dependent biotransformations with native microorganisms on an industrial scale (see [24]). However, as the substrates of these latter reactions are not steroids, they fall out of the range of this survey. 6.3.1 11a-Hydroxylation

In the early 1950s it was found that progesterone is hydroxylated in the 11αposition by Rhizopus arrhizus and R. nigricans [25] and by Aspergillus niger [26]. This microbial reaction was an important breakthrough in partial synthesis of glucocorticoids because it allowed for the first time the functionalization of the non-activated C-11. The 11α-hydroxysteroids are of interest as intermediates in some cases, but the chemical transformation of the 11α-hydroxy group into the desired 11β-configuration is also possible by chemical means [23]. Biochemically, the best characterized are the steroid 11α-hydroxylase systems from the two Rhizopus species. In the 1970s, Breskvar and Hudnik-Plevnik showed that the 11α-hydroxylase of R. nigricans is a cytochrome P450 enzyme by demonstrating the characteristic CO-difference spectrum with a maximum at 450 nm as well as the inhibtion of the enzyme by carbon monoxide; and this enzyme, together with the corresponding reductase, was subsequently shown to be microsomally located [27–29]. While in non-induced microsomal preparations the P450 content is very low (100 m3 fermentation scale with volumetric productivities higher than 1 g L −1 h−1 and diacid concentrations much higher than 100 g L −1. The aerobic, viscous fermentation system to produce long-chain DCAs by Candida tropicalis may run into gas–liquid exchange limitations. Addition of hydrogen peroxide as a liquid oxygen source improved the productivities in a laboratory system by 25%. A final concentration of about 154 g L −1 DCA was obtained [20]. Other different systems have been described, including the biotransformation to DCA by immobilized Cryptococcus cells [21], biocatalyst improvement by cell treatment with organic solvents and detergents [22], and other DCA producers such as Candida cloacae for the oxidation of long-chain fatty acids to dioic acids [23]. Recombinant Candida tropicalis strains may also transform other substrates as long as they show a linear alkyl functionality in the molecules (Fig. 8.4).

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Fig. 8.4 Possible feedstocks and products for bioconversions with Candida tropicalis.

8.3 Aromatic Compounds/Fine Chemicals

Selective microbial oxidations are well established in industrial biotransformations for fine chemicals to produce valuable chemical intermediates or active pharmaceutical ingredients (APIs) [3, 24]. The following examples focus on the selective hydroxylation of aromatic compounds. An example for the oxidation of an aliphatic side-chain to an aldehyde and the epoxidation of styrene will be discussed as well. 8.3.1 Conversion of Toxic Compounds: Catechols

Catechols substituted at the 3-position are important starting materials for pharmaceutical compounds such as L-DOPA analogs. The hydroxylation of 2phenylphenol to 3-phenylcatechol in an aqueous medium with suspended whole cells of Escherichia coli JM 101 has been successfully demonstrated up to a 300 L scale with a conversion rate of 97%, a selectivity of 85%, and a yield of 83% [25] (Fig. 8.5). Due to the toxicity of the educts and products the process is run with an in situ recovery approach. The substrate 2-phenylphenol is fed continuously at 0.45 g L −1 h−1, which is below the toxic level for the cells to be oxidized to the product 3-phenylcatechol. After complete substrate conversion the whole reaction mixture including cells passes a solid hydrophobic polystyrene based resin in an external fluidized bed and back to the main reactor. Finally the product

8.3 Aromatic Compounds/Fine Chemicals

Fig. 8.5 Bioconversion of 2-phenylphenol to 3-phenyl catechol in aqueous medium with suspended whole cells.

Fig. 8.6 Selective microbial oxidation of 2-phenoxypropionic acid (POPS) with suspended cells of Beauveria bassica in aqueous media to (R)-2-(4-hydroxyphenoxy)propionic acid (HPOPS).

can be recovered by methanol elution and purified by recrystallization in n-hexane. 8.3.2 Production of (R)-2-(4-Hydroxyphenoxy)propionic Acid

(R)-2-(4-Hydroxyphenoxy)propionic acid (HPOPS) is used as an intermediate for enantiomerically pure compounds for the production of herbicides of the aryloxyphenoxy propionic type. Products with a chemical purity >99.5% can be achieved by using a selective oxidation of 2-phenoxypropionic acid (POPS) with suspended whole cells of Beauveria bassica in aqueous media. In a 120 m3 continuous stirred reactor 7 g L −1 h−1 can be achieved [26] (Fig. 8.6). The oxidase involved in these reactions recognizes structural elements of an aromatic ring system and carboxylic acid. The para-position is preferred. Enantiomeric excess up to 96% can be obtained. The oxidase has a very broad substrate spectrum, producing compounds such as 2-(4-hydroxy-phenoxy)-butyric acid, 4(4-hydroxy-phenoxy)-benzoic acid, 2-(2-fluoro-4-hydroxy-phenoxy)-propionic acid, 2-(6-hydroxy-naphtalen-2-yloxy)-propionic acid, or (4-hydroxy-naphtalen-1-yloxy)acetic acid [24].

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Fig. 8.7 Selective oxidation of pseudocumene in a liquid two-phase system by recombinant E. coli to 3,4-dimethylbenzaldehyde (DMB aldehyde).

8.3.3 Selective Oxidation to Aromatic Aldehydes with Recombinant Cells [27]

3,4-Dimethyl-benzaldehyde can be exclusively produced in a complex multistep oxidation process at the laboratory scale. The recombinant whole cell catalysis in a two-liquid phase system was carried out with pseudocumene as substrate. Suitable concentrations of the substrate or the intermediate 3,4-dimethylbenzyl alcohol inhibited the further oxidation to 3,4-dimethylbenzoic acid. The recombinant E. coli expressed the xylene monooxygenase of Pseudomonas putida mt−. This kinetically controlled process demonstrated its industrial feasibility in 30 L fermentation working volume with a productivity of 31 g L −1 day−1, a product concentration of 37 g L −1, a purity of 97%, and an overall yield of 65% 3,4-dimethyl-benzaldehyde (Fig. 8.7). 8.3.4 Styrene Oxide Production in a Two-Liquid Phase System [2]

A whole cell biotransformation in a liquid-two phase system with toxic styrene and toxic transformation product styrene oxide was run in a two-liquid phase 30 L fed-batch bioconversion to produce 384 g styrene oxide with an enantiomeric excess of >97% after final distillation. The biotransformation reaction with recombinant E. coli JM101 (pSPZ10) expressing styrene monooxygenase of a Pseudomonas sp. strain VLB120 itself is highly selective in the reaction with an enantiomeric excess >99% (Fig. 8.8).

8.4 Heterocyclic Compounds

A series of biocatalytic routes for the production of functionalized N-heterocycles has been developed by chemical and pharmaceutical companies in cases where chemical synthesis is not efficient. The examples demonstrate how selec-


Fig. 8.8 Whole cell biotransformation of styrene to styrene oxide in a two-liquid phase system by recombinant E. coli JM101.

Fig. 8.9 Selective oxidation of the methyl group of 2,5dimethylpyrazine to 5-methylpyrazine-2-carboxylic acid without further degradation by P. putida ATCC 33015.

tive reactions of the ring hydroxylation of aromatic N-heterocycles can be controlled after selecting the suitable biocatayst. The selectivity is controlled either by inducing the enzymes specific for the reaction or by inhibiting undesired sidereactions [3]. 8.4.1 Enzymatic Oxidation of Methyl Groups in Aromatic Heterocycles

P. putida ATCC 33015, previously grown on p-xylene, is capable of oxidizing many methylated five- and six-membered rings to the corresponding monocarboxylic acids without decarboxylation or ring hydroxylation. The performance of the oxidation system was studied in greater detail with 2,5-dimethylpyrazine (Fig. 8.9) [28]. The oxidation product 5-methylpyrazine-2-carboxylic acid is an intermediate

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for the production of an antilipolytic drug. High product concentrations and high yields were obtained with growing cells. p-Xylene 75% (by volume) as growth substrate and 25% 2,5-dimethylpyrazine were added in large-scale fermentations. After 54 h, concentrations up to 20 g L −1 of 5-methyl-2-pyrazinecarboxylic acid were achieved (Fig. 8.9). 8.4.2 Preparation of 6-Hydroxynicotinic Acid

6-Hydroxynicotinic acid (Fig. 8.10), a derivative of niacin, serves as a versatile building block predominantly in the synthesis of modern insecticides. Achromobacter xylosoxidans LK1 grows on niacin and is capable of accumulating up to 65 g L −1 6-hydroxynicotinic acid with an overall yield of 90% [29]. At niacin concentrations higher than 10 g L −1 h−1 the second enzyme of the catabolic pathway (6-hydroxynicotinate hydroxylase) is strongly inhibited (Fig. 8.10). 8.4.3 Preparation of 5-Hydroxypyrazinecarboxylic Acid

A new two-step biotransformation in whole cells of Agrobacterium DSM 6336 was developed for the preparation of 5-hydroxypyrazinecarboxylic acid (Fig. 8.11) from the starting material 2-cyanopyrazine [30]. The product is used as an intermediate for novel antituberculosis drugs. The nitrile group is hydrolyzed to pyrazinecarboxylic acid by whole cell catalysis. The following regioselective hydroxylation to 5-hydroxypyrazinecarboxylic acid yielded up to 40 g L −1. The required enzymes have to be induced by growing Agrobacterium sp. DSM 6336 on 3-cyanopyridine (Fig. 8.11). 8.4.4 Preparation of 6-Hydroxy-(S)-nicotine and 4-[6-Hydroxypyridin-3-yl]4-oxobutyrate

(S)-Nicotine was used as a starting material for the biocatalytic production of functionalized pyridines used as precursors for drug production. With cells from Arthrobacter oxydans NRRL-B-3603 and Pseudomonas sp. DSM 8653 it was possible to produce up to 30 g L −1 6-hydroxy-(S)-nicotine with an isolated yield of 51% and up to 15 g L −1 of 4-[6-hydroxypyridin-3-yl]4-oxobutyrate [31] (Fig. 8.12).

Fig. 8.10 Bioconversion of niacin to 6-hydroxynicotinic acid by Achromobacter xylosoxidans LK1.


Fig. 8.11 Two-step biotransformation with whole cells of Agrobacterium DSM 6336 to produce 5hydroxypyrazinecarboxylic acid from 2-cyanopyrazine.

Fig. 8.12 Preparation of 6-hydroxy-(S)-nicotine and 4-[6hydroxypyridin-3-yl]4-oxobutyrate from plant nicotine.

8.4.5 Bulk Chemicals/Indigo

One of the world’s most important textile dyes is indigo, which is used for blue jeans coloring. The market is more than 12 000 tons per year with prices of approximately 20 euros per kg. The 250 million euro indigo market corresponds to the dying of more than 1 billion pairs of jeans.

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Today, indigo is synthesized chemically. For over 30 years anthranilic acid was used by major manufacturers, such as BASF and Hoechst. Currently aniline, formaldehyde, and hydrogen cyanide are converted to phenylglycine which reacts to indoxyl by sodamide under alkali catalysis at approximately 200–300 ºC [32]. In recent years, the synthetic process using highly toxic chemicals has been discussed critically with respect to environmental aspects. Indigo is clearly one target for the substitution of chemical synthesis by biotechnological approaches. Starting from indole, there are principally two ways to synthesize the reactive intermediate indoxyl, which converts spontanously to indigo in the presence of oxygen by water elimination [33]. The first is via cisindol-2,3-dihydrodiol (naphtaline-dioxygenase) and spontaneous decomposition to indoxyl. The second is direct hydroxylation of indole by a xylene oxidase in whole cell systems (Fig. 8.13). Indole can be generated from tryptophan by tryptophanase catalysis. Genencor [34] have developed a bacterial system capable of producing indigo on commercial scale by feeding glucose to a genetically modified E. coli strain. This strain is able to produce indole in large quantities as a precursor for the reaction to indoxyl. Growing cells induce the naphthalene dioxygenase enzyme, converting cellular indole to the unstable cis-indole-2,3-dihydroindole. This intermediate reacts under water elimination to indoxyl and in a next oxidation step to indigo. Genes from different sources might be used in E. coli as host, for example from Pseudomonas putida [36, 37] encoding naphthalene dioxygenase. To use glucose as substrate in the fermentation process the tryptophan pathway has been modified to build up high levels of cellular indole. Severe inactivation of the first enzyme of aromatic biosynthesis by the spontaneous chemical conversion of indoxyl to indigo has been observed [38]. These types of processes offer big advantages from the environmental point of view. Engineered strains were used in fermenters up to 50 000 L [35]. However, the technology is quite expensive and production costs are prohibitive at present. Many developments still need further improvement to be realized on large scale. The second approach – using recombinant bacterial systems hosting hydroxylase genes – is currently under development for the commercial production of indigo. For example, an E. coli system carrying the multicomponent phenol hydroxylase genes from Acinetobacter calcoaceticus produced low amounts of indigo (52 mg L −1) from indole in the presence of diphenylmethane. The donor strain Acinetobacter sp. ST-550 grown in a two-phase system also produces indigo from indole in the presence of a large volume of diphenylmethane and at a high level of indole [37]. Enzyme engineering is an additional powerful tool allowing the controlled use of selective microbial oxidations to produce different indigo-like dyes [39]. Genetic modifications of the single enzyme toluene ortho-monooxygenase (TOM) resulted in a wide range of color formations derived from indole as substrate. Colonies expressing these altered TOM enzymes developed by saturation mutagenesis ranged in color from blue, green, and purple to orange. The specificity


Fig. 8.13 Pathways to indigo.

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of TOM mutants expressed in E. coli TG1 for the indole hydroxylation was studied. Derived from the single TOM template, enzymes were identified that produced different types of oxidation products such as isoindigo, indigo, indirubin, and isatin.

8.5 Glycerol Conversion to Dihydroxyacetone

Dihydroxyactone (DHA) is mainly used as a tanning active, as a chemical intermediate, or as a reducing agent [44]. The production of DHA affords the selective oxidation of the hydroxylgroup at 2-position of glycerol (Fig. 8.14). Chemical synthesis of DHA by catalytic reaction of formaldehye at high pressure and temperatures is not sufficiently selective for industrial production [40, 41] in contrast to the selective glycerol oxidation by Gluconobacter oxydans [42, 43]. DHA is produced industrially in classical batch fermentation with starting concentrations of 150–200 g L −1 glycerol [44]. The process with Gluconobacter oxydans is characterized by strong substrate and product inhibition. Thus optimization approaches are attractive to overcome these productivity limitations. For example, final product concentrations of 128 g L −1 and volumetric productivities of 4 g L −1 h−1 were described in continuous processes with Acetobacter suboxydans [45], but due to the complexity and contamination risks these approaches have not been implemented on an industrial scale. A cyclic process characterized by retaining biomass resulted in productivities of 22.8 g L −1 h−1 but yielded a relatively low product concentration of 60 g L −1 due to product inhibition [46]. New interesting approaches with Gluconobacter suboxydans demonstrate the productivity improvement potential of a cyclic two-step fed-bach process. A viable, not irreversibly product-inhibited culture was maintained in the first reactor stage up to a predetermined DHA threshold value in the range 100–110 g L −1. In the second reactor stage, high final product concentrations of up to 220 g L −1 were reached, while the culture was irreversibly product-inhibited [47, 48].

Fig. 8.14 Glycerol conversion to dihydroxyacetone (DHA).

References

8.6 Perspectives

Selective microbial oxidations have been implemented successfully in many industrial processes and provide enormous opportunities for the future. White biotechnology is regarded as a driver of sustainable growth, especially in Europe and the USA, and will rely on these production methods. A variety of tools have been devoloped and will be further developed in biochemical engineering and recombinant DNA technology to optimize the biocatalyst and the process set-up. Among others, raw material suppliers such as ADM or Cargill, chemical companies such as BASF, DSM, Lonza, Dupont, and DowChemicals, and biocatalyst suppliers such as Novozymes, Genencor, Diversa, or Maxygen are investing in these technologies to be prepared for the next decade.

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8 Selective Microbial Oxidations in Industry 14 Cao, Z., Gao, H., Liu M., Jiao, P. Engineering the acetyl-CoA transportation system of Candida tropicalis enhances the production of dicarboxylic acid. Biotechnol J 2006, 1, 68–74. 15 Uemura, N., Taoka, A., Takagi, M. Production of dicarboxylic acids by fermentation. World Conference on Biotechnology of the Fats and Oils Industry 1988, 148–152. 16 Liu, S.-C., Li, C., Fang, X., Cao, Z. Optimal pH control strategy for highlevel production of long-chain α,ωdicarboxylic acid by Candida tropicalis. Enzyme Microb Technol 2004, 34, 73–77. 17 Craft, D.L., Madduri, K.M., Eshoo, M., Wilson, C.R. Identification and characterization of the CYP52 family of Candida tropicalis ATCC 20336, important for the conversion of fatty acids and alkanes to α,ω-dicarboxylic acids. Appl Environ Microbiol 2003, 69, 5983–5991. 18 Eirich, L.D., Craft, D.L., Steinberg, L., Asif, A., Eschenfeldt, W.U., Stols, L., Donnelly, M.I., Wilson, C.R. Cloning and characterization of three fatty alcohol oxidase genes from Candida tropicalis strain ATCC 20336. Appl Environ Microbiol 2004, 70, 4872–4879. 19 Picataggio, S., Rohrer, T., Deanda, K., Lanning, D., Reynolds, R., Mielenz, J., Eirich, L.D. Metabolic engineering of Candida tropicalis for the production of long-chain dicarboxylic acids. Bio/Technology 1992, 10, 894–898. 20 Jiao, P., Huang, Y., Li, S., Hua, Y., Cao, Z. Optimal pH control strategy for highlevel production of long-chain α,ωdicarboxylic acid by Candida tropicalis. Biotechnol Bioeng 2001, 75, 456–462. 21 Chan, E.-C., Kuo, J. Biotransformation of dicarboxylic acid by immobilized Cryptococcus cells. Enzyme Microb Technol 1997, 20, 585–589. 22 Chan, E.-C., Kuo, J., Lin, H.-P., Mou, D.-G. Stimulation of n-alkane conversion to dicarboxylic acid by organic-solventand detergent-treated microbes. Appl Microbiol Biotechnol 1991, 34, 772–777. 23 Green, K.D., Turner, M.K., Woodley, J.M. Candida cloacae oxidation of long-

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chain fatty acids to dioic acids. Enzyme Microb Technol 2000, 27, 205–211. Liese, A., Seelbach, K., Wandrey, C. Industrial Biotransformations. Weinheim: Wiley-VCH Verlag, 2000. Held, M., Schmid, A., Kohler, H-P.E., Suske, W., Witholt, B., Wubbolts, M.G. An integrated process for the production of toxic catechols from toxic phenols based on a designer biocatalyst. Biotechnol Bioeng 1999, 62, 641–648. Dingler, C., Ladner, W., Krei, G., Cooper, B., Hauer, B. Preparation of (R)-2-(4hydroxyphenoxy)propionic acid by biotransformation. Pesticide Sci 1996, 46, 33–35. Bühler, B., Bollhalder, I., Hauer, B., Witholt, B., Schmid, A. Chemical biotechnology for the specific oxyfunctionalization of hydrocarbons on a technical scale. Biotechnol Bioeng 2003, 82, 833–842. Kiener, A. Enzymatic oxidation of methyl groups on aromatic heterocycles: a versatile method for the preparation of heteroaromatic carboxylic acids; Patent no. EP-B 0442430. Angw Chem Int Ed 1992, 31, 774–775. Kulla, H.G. Enzymatic hydroxylations in industrial application; Patent EP-B 0152948; EP-B 0152949. Chimia 1991, 45, 81–85. Wieser, M., Heinzmann, K., Kiener, A. Bioconversion of 2-cyanopyrazine to 5hydroxypyrazine-2-carboxylic acid with Agrobacterium sp. DSM 6336. Patent EP-B 0578137. Appl Microbiol Biotechnol 1997, 48, 174–176. Roduit, J.P., Wellig, A., Kiener, A. Renewable functionalized pyridines derived from microbial metabolites of the alkaloid (S)-nicotine; EP-B 0713869. Heterocycles 1997, 46, 1687–1702. Cofrancesco, A.J. Dyes, natural. KirkOthmer Encyclopedia of Chemical Technology – Online, DOI: 10.1002/047123 8961.0425051903150618.a01. 2000 Online Posting, John Wiley & Sons. Ensley, B.D., Ratzkin, B.J., Osslund, T.D. Expression of naphthalene oxidation genes in Escherichia coli results in the biosynthesis of indigo. Science 1983, 222, 167–169.

References 34 Murdock, D., Ensley, B., Serdar, C., Thalen, M. Construction of metabolic operons catalyzing the de novo biosynthesis of indigo in Escherichia coli. Bio/Technology 1993, 11, 381–386. 35 Bialy, H. Biotechnology, bioremediation, and blue genes. Nat Biotechnol 1997, 15, 110–111. 36 Buehler, B., Schmid, A., Hauer, B., Witholt, B. Xylene monooxygenase catalyzes the multistep oxygenation of toluene and pseudocumene to corresponding alcohols, aldehydes, and acids in Escherichia coli JM101. J Biol Chem 2000, 275, 10085–10092. 37 Doukyu, N., Toyoda, K., Aono, R. Indigo production by Escherichia coli carrying the phenol hydroxylase gene from Acinetobacter sp. strain ST-550 in a water-organic solvent two-phase system. Appl Microbiol Biotechnol 2003, 60, 720–725. 38 Berry, A., Dodge, T.C., Pepsin, M., Weyler, W. Application of metabolic engineering to improve both the production and use of biotech indigo. J Ind Microbiol Biotechnol 2002, 28, 127–133. 39 Rui, L., Reardon, K.F., Wood, T.K. Protein engineering of toluene orthomonooxygenase of Burkholderia cepacia G4 for regiospecific hydroxylation of indole to form various indigoid compounds. Appl Microbiol Biotechnol 2005, 66, 422–429. 40 Gehrer, E., Harder, W., Knuth, B., Vogel, H. Patent DE 4122669. 1993.

41 Ohrem, H.L., Westmeier F. Patent US 5770411. 1995. 42 Gupta, A., Gupta, K., Singh, K. Gluconobacter oxydans: Its biotechnological applications. J Mol Microbiol Biotechnol 2001, 3, 445–456. 43 Ohrem, H.L., Voss, H. Process model of the oxidation of glycerol with Gluconobacter oxydans. Proc Biochem 1996, 66, 295–301. 44 Falbe, J., Regitz, M. (eds) Roempps Chemie Lexikon, Vol. 2. Stuttgart: Franck, 1997. 45 Izuo, N., Nabe, K., Yamada, S., Chibata, I. Production of dihydroxyacetone by continuous cultivation of Acetobacter suboxydans. J Ferment Technol 1980, 58, 221–226. 46 Hekmat, D., Bauer, R., Fricke, J. Optimization of the microbial synthesis of dihydroxyacetone from glycerol with Gluconobacter oxydans. Bioprocess Biosyst Eng 2003, 26, 109–116. 47 Bauer, R., Katsikis, N., Varga, S. Hekmat, D. Study of the inhibitory effect of the product dihydroxyacetone on Gluconobacter oxydans in a semicontinuous two-stage repeated-fed-batch process. Bioprocess Biosyst Eng 2005, 28, 37–43. 48 Bauer, R., Hekmat, D. Development of a transient segregated mathematical model of the semicontinuous microbial production process of dihydroxyacetone. Biotechnol Progr 2006, 22, 278–284.

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9 Preparation of Drug Metabolites using Fungal and Bacterial Strains Oreste Ghisalba and Matthias Kittelmann

9.1 Introduction

Within the drug discovery and development process the investigation of the metabolic fate of drug candidates (phase I and phase II metabolism) is a crucial issue and of utmost importance. As a general trend in all big pharmaceutical companies (as well as in small and medium sized companies dedicated to drug discovery) the time span allotted for drug discovery and development activities has been very significantly shortened during the past two decades. This change was triggered by economic pressure and to a large extent enabled by the introduction of novel screening techniques based on biologically validated drug targets in combination with high performance (bio)analytical and (bio)synthetic methods. As a consequence of this reduced R&D time frame, access to metabolic information on potential drug candidates is needed much earlier than in the past. This requires an advancement of the metabolism-related investigations from the early development phases to the late discovery phases, leading to very significantly increased numbers of compounds to be tested and numbers of metabolites to be prepared for biological evaluation. Conventional drug metabolism studies used to rely on the use of small animal models and in vitro assays. However, there are significant differences in the organ distribution, the regulation, and the activities/selectivities of the drugmetabolizing enzymes (DMEs) between mammalian species. Such variations can lead to the identification of metabolites that are not necessarily the same as those observed in human systems. This has led to a shift – at least partially – in emphasis away from in vivo experiments with animals to in vitro studies with human DMEs in order to increase the accuracy of prediction of the metabolic fate of the drug candidates. Existing in vitro assay systems for analytical applications are based on human liver slices, human hepatocytes, microsomal preparations, or liver homogenates. The choice is dictated by the goals.

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9 Preparation of Drug Metabolites using Fungal and Bacterial Strains

The DMEs are a diverse group of enzymes that are responsible for metabolizing a vast array of xenobiotic compounds including drugs, environmental pollutants, and endogenous compounds such as steroids and prostaglandins [1, 2]. From an enzymological point of view, they are most noted for their broad substrate specificity: some members of the cytochrome P450 and flavin monooxygenase families are known to metabolize more than 50 structurally diverse compounds. Conceptually, the DMEs have been divided into two groups: • The phase I enzymes catalyze (mainly oxidative) reactions that generally result in the introduction of a functional group into substrate molecules. The phase I enzyme families include cytochromes P450 (P450s or CYPs) and flavin-dependent monooxygenases (FMOs), but also monoamine oxidases (MAOs), dehydrogenases, isomerases, hydrolases, and others. • The phase II enzymes catalyze conjugative reactions that usually result in the formation of soluble compounds that are more readily excreted. The phase II enzyme families include the UDP-glucuronosyl transferases (UGTs), glutathione transferases (GSTs), sulfotransferases (SULTs), and N-acetyltransferases (NATs). The use of recombinant human DMEs (CYPs, UGTs, and SULTs) has already played an important role in metabolism-related R&D efforts, but is a field that is still in its early stages. Whereas analytical applications are straightforward and technically relatively simple, the preparative application of such systems for the synthesis of multi-milligram to multi-100 mg quantities of metabolites proved to be a challenging task as not only the reaction engineering but also the isolation and purification of the individual metabolites from complex metabolite mixtures can be very difficult and time consuming. Microbial equivalents of human DMEs, especially for CYP-type reactions are an additional asset especially of interest when larger amounts of metabolites must be prepared. At Novartis, standardized procedures for the preparation of the biocatalysts, for bioconversion (phase I and phase II metabolic reactions), and for product isolation have now been successfully established but can still be improved based on the growing application experience with the systems. As the focus of this review is on biological oxidation, the phase II reactions will not be further discussed.

9.2 Phase I Drug-Metabolizing Enzymes

Biotransformation phase I converts foreign compounds (drugs) into more watersoluble metabolites by the addition of functional groups (-SH, -OH, -NH2,

9.2 Phase I Drug-Metabolizing Enzymes

-COOH) necessary for the phase II reactions, and by the exposure of pre-existing functional group (e.g. via esterases, amidases, etc.). The types of oxidative reactions catalyzed by P450s (FMOs) include: • aliphatic, aromatic, allylic or benzylic oxidation/ hydroxylation • epoxidation of aromatic or alkene double bonds • N-, O-, S-dealkylation • N-oxidation of secondary and tertiary amines • S-oxidation of S-alkyl derivatives • N-hydroxylation of secondary and tertiary amines. Other types of oxidative phase I reactions are: • oxidative deamination of primary amines (by monoamine oxidases) • Baeyer–Villiger reaction (with FMOs). The oxidation of organic molecules by CYPs (or FMOs) is quite complex [3], but the overall reaction can be represented simply by the following equation: RH + O2 + NADPH + H + → ROH + H2O + NADP+ In the case of CYPs, NADPH supplies electrons via the flavin domain of the associated NADPH-P450 oxidoreductase. In some cases, the second electron can come from NADH via cytochrome b5 reductase and cytochrome b5 [4]. There are at least 16 families of P450 isozymes expressed in humans. The CYP3A, 2D6, and 2C isozymes metabolize most of the drugs for which a specific isozyme has been identified [2]. FMOs, like CYPs, catalyze the oxidation of organic compounds by molecular oxygen using NADPH as a source of electrons for the reduction of one of the oxygen atoms. However, they are mechanistically distinct from the P450s in that they react with oxygen and NADPH in the absence of substrate to form a 4αhydroperoxy flavin enzyme intermediate. Thus, the FMOs exist in an activated form in the cell, and their interaction with a nucleophilic group such as an amine, thiol, or phosphate is all that is required for completion of the catalytic cycle. This capacity is a possible explanation for the extremely broad substrate specificity of the FMO isozymes. Compared to the P450s, there are relatively few FMOs (five subfamilies in humans). The FMO group of enzymes are important in the metabolism of some drugs and toxins and their relative abundance in most mammalian tissues indicates that they contribute substantially to detoxification of xenobiotics. In oxidative metabolism of xenobiotics/drugs FMOs or MAOs often generate the same metabolites as those generated by CYPs. This is especially true for Nand S-oxygenations where CYPs and FMOs can yield identical metabolites. As a consequence, the identification of the enzyme(s) responsible for a specific biotransformation can be very tricky.

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9.3 Needs and “Platforms” for the Generation of Drug Metabolites

From an industrial perspective the needs/applications listed in Table 9.1 can be defined in relation to in vitro investigation of oxidative drug metabolism and to enzymatic preparation of drug metabolites. The choice of the biocatalyst(s) is dictated by the tasks to be performed. The substrates to be investigated for relevant metabolism/metabolites can be: • Synthetic drug candidates/drugs/prodrugs • Biologically active natural products (e.g. microbial secondary metabolites) • Biologically active semi-synthetic derivatives of natural products. At Novartis Bioreactions Laboratories the in vitro DME platforms for phase I oxidative metabolism studies and preparative scale metabolite synthesis established so far are the following. Table 9.1 Types of in vitro biocatalyst applications in metabolism-related drug R&D activities.

Application

Suitable biocatalysts[a] (enzyme preparations/whole cells, etc.)

1. Analytical profi ling of drug candidates or drugs (a) Establishment of metabolic pattern with tentative structural assignments (HPLC-MS) (b) Prediction of the most favored metabolic reactions (in vivo) 2. Small-scale synthesis of human drug metabolites (milligram scale) (a) Structure elucidation/confi rmation with isolated/purified metabolites (MS, NMR) 3. Preparative scale synthesis of major human drug metabolites (a) For pharmacological and toxicological evaluation (10–100 mg scale) (b) For derivatization programs (100 mg to multi-gram scale) 4. Identification and production of new (i.e. non-human) metabolites with new or modified biological properties as lead compounds for derivatization programs (100 mg to multi-gram scale)

Commercial (human) liver preparations Recombinant human CYPs as whole cells or microsomes (Recombinant mammalian CYPs) (Microbial biocatalysts [b] , whole cells)

Recombinant human CYPs as whole cells or microsomes (Microbial biocatalysts [b] , whole cells) Recombinant human CYPs as whole cells Microbial biocatalysts [b] , whole cells

Microbial biocatalysts, whole cells

a According to Novartis experience. b With known biocatalytic similarity to mammalian/human CYPs. Biocatalysts in parentheses are less suitable.

9.4 Microbial Models for Oxidative Drug Metabolism

9.3.1 Recombinant Human Cytochrome P450 (rhCYP) Systems (acquired from British Technology Group/University of Dundee)

This set of fourteen recombinant Escherichia coli strains functionally coexpressing individual human CYP isoforms together with human NADPH-P450 reductase comprises all human CYP isoforms with proven relevance in drug metabolism (i.e. CYPs 1A1, 1A2*, 1B1, 2A6, 2B6, 2C8, 2C9*, 2C18, 2C19*, 2D6*, 2E1, 3A4*, 3A5, and 4A11). The isoforms marked with * are considered to be the major five in drug metabolism. As an analytical tool, this platform enables us not only to determine the overall phase I metabolite patterns of the investigated drug candidates but also to identify the responsible CYP isoform(s) for each metabolite. The systems have been successfully used for the synthesis of major metabolites up to multi-100 mg scale. 9.3.2 Microbial Strains Performing Oxidative Reactions (in-house technology)

These are >150 microbial strains (bacteria, fungi) performing hydroxylations, epoxidations, dealkylations, etc. This platform is complementary to the rhCYPbased technology and very useful in cases where larger quantities (multi-100 mg to multi-gram scale) of metabolites are requested. In many cases the microbial oxygenases are characterized by higher turnover numbers and better operational stabilities than the mammalian phase I DMEs. The strains were selected based on literature data and on in-house experience from screening efforts. As is the case with many microbial processes and reactions described in the literature the oxygenase enzymes (CYPs, FMOs, dioxygenases, etc.) responsible for a given biotransformation have often not been purified and characterized as the reactions were performed with whole microbial cells. Microbial (bacterial, fungal) oxidative metabolism is not always directly comparable to mammalian systems.


The focus of this review is on correlation of microbial oxidative biocatalysts with human/mammalian cytochrome P450 systems as much less knowledge, experience, and literature are available on FMOs for both mammals and microbes. Although numerous mammalian cytochromes P450 (CYPs) have been extensively studied, detailed research on their microbial counterparts for many years has been limited to the P450 systems of Pseudomonas putida and Saccharomyces cerevisiae [5]. Only in recent years have a number of other prokaryotic and eukaryotic P450 enzymes of microbial origin been studied in detail. The studies

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have emphasized the role of these enzymes – similar to their mammalian counterparts – in the oxidation of chemicals of an endobiotic and xenobiotic nature. Over the past 20 years, several reports have described systematic studies dedicated to the practical problem of producing potentially important mammalian drug metabolites using microorganisms. There are basically two major types of P450 systems known so far (in aerobic organisms). The first is the system found in the endoplasmic reticulum of eukaryotic microorganisms (type I, Fig. 9.1), which consists of an FAD- and FMN-containing reductase for transport of electrons from NADPH to the P450 component. It appears that one reductase can transfer electrons to more than one P450 component in mammalian systems. The microsomal P450 enzymes exhibit broad and overlapping catalytic activities and are perhaps the most non-specific enzymes known. The recombinant E. coli strains coexpressing human CYP and human P450 reductase mimic this arrangement. In contrast, the second type of P450 system (type II, Fig. 9.2), which is found in mitochondria and bacteria, generally possesses very tight substrate specificities. Type II P450 systems, in addition to an FAD-containing reductase (ferredoxin reductase), use a small redox iron–sulfur protein (ferredoxin) for transfer of electrons from NAD(P)H to the P450 component. A third type of organization of P450 components has been reported for Bacillus megaterium. In this microorganism a catalytically self-sufficient P450, which does not require the presence of ancillary electron transport proteins, performs its oxidative reactions in the presence of the reduced pyridine nucleotide [6]. For many decades, authors have proposed the use of microbial metabolic systems to predict the possible fate of drugs in mammals or humans. Pioneers

Fig. 9.1 Scheme of type I cytochrome P450 system.

Fig. 9.2 Scheme of type II cytochrome P450 system.


periodically delivering attempts to correlate mammalian and microbial metabolic data include the research groups of Robert Azerad (CNRS, University René Descartes, Paris), F. Peter Guengerich (Vanderbilt University, Nashville, Tennessee), and Robert V. Smith (University of Texas, Austin)/John P. Rosazza (University of Iowa, Iowa City). However, this option is to be regarded and exploited with caution. In view of the arsenal of investigation methods available today (see above) it is much safer to start a drug metabolism study with an analytical profi ling step using a human (mammalian) DME preparation (tissue fraction or recombinant enzyme) as outlined in Table 9.1 and then – in a second step, if needed – search for a microbial biocatalyst once the pharmacologically relevant metabolic pattern is known and larger quantities of defined major (or minor) metabolites are requested for biological evaluation. An in-depth literature analysis and also our in-house laboratory experience show that for a given drug candidate (substrate) the metabolic patterns generated by mammalian/human DME systems can be very different from the microbial ones. Several situations can occur in comparative metabolism studies (HPLC-MS analysis): 1. No microbial (i.e. bacterial or fungal) metabolism is found even after extensive screening efforts. In this case it can be concluded that the envisaged metabolism/metabolites is/ are restricted to mammalian systems. 2. Microbial metabolism is found but the pattern is very different from mammalian/human metabolism because: (a) the abundance of individual metabolites is different, (b) the metabolite(s) of interest is/are not formed or only in very low amounts, or (c) the molecular weight distribution of the detected metabolites is different (i.e. new metabolites of potential interest are found that were not detected with mammalian/human DME system). 3. The microbial metabolism pattern is comparable with the previously detected mammalian/human pattern. This is the ideal case, where there is: (a) comparable distribution of metabolites, (b) comparable relative abundances or metabolites, or (c) the desired metabolites are formed in reasonable quantities to justify isolation and purification efforts. In cases 1 and 2 one has to try to synthesize the desired metabolites (e.g. with rhCYPs). In case 2 a preparative scale microbial biotransformation could be of interest to produce the new (non-human) metabolite(s) for further evaluation. In case 3 a preparative scale microbial biotransformation can give access to larger amounts of the desired metabolites. In some cases an additional reaction engineering effort is required to enhance the yield of the target metabolite(s).

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9.4.1 Prokaryotic P450s

Prokaryotic P450 enzymes are generally multicomponent soluble systems that fall within the type II category. The majority of these enzymes are either extremely substrate specific or act only on a limited number of structurally related substrates. Most of the enzymes are inducible. Prokaryotic P450 monooxygenases resemble the steroidogenic P450 systems found in adrenal mitochondria. A selection of oxidative biotransformation examples with prokaryotic systems is compiled in Table 9.2. The P450soy (from Streptomyces griseus) is a very rare case, where a prokaryotic P450 resembles its mammalian counterparts in its broad substrate specificity and also in its ability to activate promutagenic chemicals. 9.4.2 Microbial Eukaryotic P450s

Microbial eukaryotic enzymes generally fall within the type I category with respect to the organization of their redox components. The particulate and unstable nature of microbial eukaryotic cytochromes P450 renders them difficult to isolate and purify. The most thoroughly studied microbial eukaryotic P450s are the enzymes from Saccharomyces cerevisiae and Candida albicans. Most of the enzymes are inducible. In comparison to prokaryotic systems, yeast and fungi possess monooxygenase systems more similar to those found in mammalian hepatic microsomes. A selection of oxidative biotransformation examples with microbial eukaryotic systems is shown in Table 9.3. Some of the listed P450s of microbial origin can mimic mammalian P450s with respect to their broad substrate range. Good candidates are the prokaryotic P450soy from S. griseus, the eukaryotic P45014DM from S. cerevisiae, and the eukaryotic P450 from Cunninghamella bainieri. These deserve further evaluation for the development of larger scale bioreactor applications for the production of metabolites. Compared with mammalian systems, the use of suitable microbial systems would have significant economic advantages. An additional option is to systematically screen a large range of biologically diverse microbes for additional P450s with broad substrate ranges in order to complement the arsenal of easier-to-produce suitable biocatalysts. The goal would be to set up a routine screening of a carefully selected group of perhaps 30–50 of “highly competent” microorganisms to quickly assess the probability of producing drug metabolites or natural product derivatives. Very early reports [12–14] list selected broadly applicable microorganisms (mainly fungi) as microbial models of mammalian aromatic hydroxylation: Aspergillus niger ATTC 9142, Aspergillus ochraceus ATCC 1008, Cunninghamella bainieri ATCC 9244, Cunninghamella blakesleeana ATCC 8688a, Curvularia lunata NRRL 2178, Gliocladium deliquescens 1086, Helicostylum piriforme QM 6945, Penicillium chrysogenum ATCC 10002, Rhizopus stolonifer NRRL 1477, Streptomyces

P450SU 1 P450SU 2 P450 soy

P450 sca-1 P450 sca-2

P450 cam

P450PM-3 P450 meg

P450PM-1

P450 system

O-Demethylation of lignin-related compounds (veratrol, guaiacol) Cholesterol oxidation Oxidation of sulfonylurea herbicides (chlor-sulfon, sulfometuron methyl, chlorimuron ethyl Oxidation of various structurally unrelated xenobiotics → aromatic hydroxylation, alicyclic hydroxylation, benzylic oxidation, O-dealkylation, non-aromatic double bond epoxidation, N-acetylation Broad substrate range Generation of mammalian metabolites of 10,11-dihydro-5H-dibenzo[a,d] cycloheptane-5-carboxamide N-Demethylation of amitriptyline, nortryptiline, codeine, and metazocine N-Demethylation of diazepam N-Demethylation of codeine Hydroxylation of acronycine to 9-hydroxy-acronycin [a] 18- and 19- hydroxylation of PGE2α and PGF2α[a] Hydroxylation of cycloalkanes and cycloalkenes to cycloalkanols and cycloalkenols

Oxidation of long chain n-alkanes Hydroxylation of danazol metabolites [a] Hydroxylation of long-chain fatty acids and their corresponding amides and alcohols Hydroxylation of long-chain fatty acids 15β-Hydroxylation of 3-oxo-Δ4 steroids n-Octane oxidation Dealkylation of p-nitroanisole Side-chain hydroxylation of guaiacol to generate catechol and formaldehyde Conversion of thromboxane B2 to dinor-tromboxane B2 [a] Dealkylation of 4-alkoxybenzoates Hydroxylation of fatty acids and hydrocarbons Oxidation of camphor, norcamphor and 1-methylnorcamphor Dehalogenation of chloropicrin and various polyhalomethanes Hydroxylation of 6-deoxyerythronolide B to erythronolide B Hydroxylation of compactin to pravastatin

Catalyzed reaction/substrate range

a These mammalian metabolites were generated by microbial biotransformations on preparative scale (up to multi-gram).

Streptomyces lincolnensis Streptomyces rimosus Streptomyces paucisporogenes Streptomyces spectabilicus Streptomyces sp. Xanthobacter sp.

ATCC 25466 NCIB 8229 ATCC 12596 NRRL 2494 UC 5761

ATCC 13273

Streptomyces griseus

Streptomyces lavendulae

ATCC 11796

Streptomyces setonii Streptomyces sp. Streptomyces griseolus

Saccharopolyspora erythreae Streptomyces carbophilus

ATCC 17453

UC 6176

ATCC 14581 ATCC 13368

ATCC 6946 ATCC 14581

Acinetobacter calcoaceticus Arthrobacter simplex Bacillus megaterium

Bacillus megaterium Bacillus megaterium Corynebacterium sp. Escherichia coli Moraxella sp. Mycobacterium rhodochrous Nocardia (NH1 strain) Pseudomonas oleovorans Pseudomonas putida

Strain collection

Microorganism

Table 9.2 Prokaryotic P450s and catalysed oxidations with free enzymes or whole cells; selection [5, 7–11].

9.4 Microbial Models for Oxidative Drug Metabolism 219

FO 6226 CBS 38448 ATCC 6227b ATCC 12939

CBS 18435 NRRL 1570

CMI 53585 NRRL 3655 IMI 199844 ATCC 9245 ATCC 36112

P45014DM

P450/P455

Three isozymes

P45014DM P450alk

P450 system

Alkane oxidation Hydroxylation of acronycine to 9-hydroxy-acronycin [a] Hydroxylation of ellipticine to 8-hydroxy-and 9-hydroxyellipticines, [a] 6-O-demethylation of papaverine [a] C11-Hydroxylation of progesterone 14α-Demethylation of lanosterol ω-Oxidation of long-chain fatty acids (lauric acid) and hydrocarbons; oxidation of hexobarbital, aminopyrine, benzphetamine, ethylmorphine, 4′hydroxylation of biphenyl Relatively broad substrate range Hydroxylation of n-alkanes Aromatic hydroxylation, N- and O-demethylation of various xenobiotics (amitriptyline, chloropromazine, codeine, diazepam, and metazocine) Broad substrate range N-Demethylation of codeine, diazepam, and metazoxine Hydroxylation of acronycine to 9-hydroxy-acronycin [a]/N-demethylation of amitriptyline, chloropromazine, codeine, diazepam, and metazocine Hydrocarbon oxidation (naphthalene), O-demethylation of 10,11dimethoxyaporphine to isoapocodeine [a] ω-1-, ω-2-, ω-3-Oxidation of fatty acid (e.g. lauric acid) Hydroxylation of danazol metabolites [a] 18- and 19-hydroxylation of 9-oxo-13-(cis)-prostenoic acid [a] Demethylation of phytoalexins (isatin) 4′-trans hydroxylation of 6-chloro-5-cyclohexylindan-1-carboxylic acid [a] Oxidation of m-cresol, patulin biosynthesis 11α-Hydroxylation of progesterone Hydroxylation of danazol metabolites [a] 14α-Demethylation of lanosterol, oxidation of benzo[a]pyrene, β-naphthylamine, cyclo-phosphamide, hexobarbital, aminopyrine, benzphetamine, ethylmorphine, etc. Broad substrate range Demethylation of aminopyrine

Catalyzed reaction/substrate range

a These mammalian metabolites were generated by microbial biotransformations on preparative scale (up to multi-gram).

Trichosporon cutaneum

Fusarium oxysporum Fusarium reticulatum Microascus trigonosporus Nectria haematococca Penicillium concavo-rugulosum Penicillium patulum Rhizopus nigricans Rhizopus stolonifer Saccharomyces cerevisiae

Cunninghamella elegans

Cunninghamella blakesleeana Cunninghamella echinulata

Candida maltosa Cunninghamella bainieri (C. echinulata) ATCC 9244

QM 1915 NRRL 315

Acremonium sp. Aspergillus alleaceus Aspergillus alleaceus

Aspergillus ochraceus Candida albicans Candida tropicalis (various strains)

Strain collection

Microorganism

Table 9.3 Microbial eukaryotic P450s and catalyzed oxidations with free enzymes/microsomes or whole cells; selection [5, 7–11]

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9.6 Correlation of Microbial Reactions with Human CYP Isozyme-Specific Reactions

rimosus ATCC 23955, and Streptomyces sp. 1158w. The hydroxylation patterns of these strains were elucidated using a broad range of aromatic test substrates (for detailed listings see [12, 13]). A later reference [10] states that the following additional collection strains are extremely useful as versatile biocatalysts: Arthrobacter sp. ATCC 19140, Beauveria bassiana ATCC 7159, Cunninghamella elegans ATCC 9245/ATCC 36112, Cunninghamella echinulata ATCC 9244/NRRL 3655, Nocardia corallina ATCC 19070, and Streptomyces rimosus NRRL 2234. Conditions and strategies for the induction of microbial P450 monooxygenases can be found in [9].

9.5 Correlation of Microbial and Mammalian Oxidative Drug Metabolism

A significant number of pharmacologically active drugs have so far been subjected to comparative metabolic studies: mammalian metabolism vs. microbial metabolism, with involvement of DMEs (mainly P450s). A very useful compilation of such data was published by Azerad [10] and is summarized in Table 9.4. The selected data in Table 9.4 clearly indicate that many types of mammalian metabolites can also be successfully generated with microbial systems. However, it must be borne in mind that in a significant number of cases specific mammalian metabolites cannot be generated with microbial systems and in other cases the microbes produce additional types of metabolites that are not found with the mammalian systems.

9.6 Correlation of Microbial Reactions with Human CYP Isozyme-Specific Reactions

A challenge for the future is to directly correlate bacterial and fungal cytochrome P450 reactions with specific human CYP isoforms performing the same reactions with a given drug. Such a correlation would hopefully facilitate strain selection and allow a straightforward shift from biotransformations with, for example, rhCYPs (analytical scale, preparative scale-up to gram quantities) to biotransformations with microbial systems for scaling-up purposes (multi-gram to large scale). In the Novartis Bioreactions Laboratories limited efforts have been made to establish such a tentative correlation between microbial CYP biocatalysts with the most important human CYP isozymes using standard model drugs (Parel, S., Kittelmann, M., Ghisalba O., Novartis unpublished results). The results of this exploratory work are listed in Table 9.5. The screening results obtained with the three human CYP1A2 substrates caffeine, phenacetin, and theophylline clearly indicate that a positive result in an investigated microbial bioconversion reaction with respect to a given human CYP isozyme substrate is not necessarily an indication of an “identical” substrate

221

C-5- and C-6 Hydroxylations 10,11-Epoxidation C-10 Hydroxylation N-Demethylation C-1 Hydroxylation N-Demethylation, 10,11-epoxidation, aromatic hydroxylations at C-1, C-2, or C-3, N-oxidation N-Demethylation and C-3 hydroxylation, C-4′ aromatic hydroxylation Terminal methyl hydroxylation and oxidation to carboxylic acid C-9 Hydroxylation N-Dealkylation Stereoinversion (R to S), C-2′ hydroxylation Allylic methyl group hydroxylation and oxidation to carboxylic acid N-Demethylation Aromatic methyl group hydroxylation Oxidative deamination, C-4′ aromatic hydroxylation C-1′ or C-3′ Hydroxylations

Bornaprine (anticholinergic) Carbamazepine (antiepileptic) 10,11-Dihydrocarbamazepine (antiepileptic)

Codeine (analgesic) Crisnatol (antitumor) Cyproheptadiene (antihistamine)

Papaverine (vasodilatator, muscle relaxant, antitumor)

Lergotrile (antiparkinson/ antiprolactin) Lucanthone (schistosomicidal) Mexiletine (antiarrythmic) MK 954 (angiotensin A II receptor antagonist)

Ellipticine (antitumor) Furosemide (diuretic) Ibuprofen (anti-inflammatory) Lapachol (antitumor antibiotic, antimalarial)

Ebastine (antihistamine)

6-O-Demethylation and 4′-O-demethylation

C-9 Hydroxylation N-Demethylation, C-4 hydroxylation, N-dealkylation N-Dealkylation, C-4-, C-5-, C-6-aromatic hydroxylations O-Deisopropylation oxidation to carboxylic acid

Acronycine (antitumor) Alosetron (5HT3 receptor antagonist) Besipirdine (cholinergic and noradrenic agonist) Bisprolol (β-blocking agent)

Diazepam (tranquilizer)

Metabolic reactions (only reaction types that were found identical for mammalian and microbial systems)[a]

Drug (pharmacological activity)

Streptomyces platensis Aspergillus sclerotiorum Cunninghamella echinulata Actinoplanes sp. Streptomyces ATCC 55293 Aspergillus alliaceus Cunninghamella echinulata

Aspergillus alliaceus Cunninghamella elegans Verticillium lecanii Beauveria bassiana

Aspergillus terreus Beauveria bassiana Cunninghamella blakesleeana

Cunninghamella echinulata, Aspergillus alliaceus Streptomyces griseus Cunninghamella elegans Cunninghamella echinulata Gliocladium eliquescens Gliocladium deliquescens Cunninghamella echinulata Streptomyces violascens Streptomyces violascens Streptomyces griseus Streptomyces griseus Cunninghamella elegans Cunninghamella elegans

Microorganisms

Table 9.4 Microbial and mammalian oxidative metabolism – comparative studies; selection of positive correlations.

222


Aspergillus fl avus Aspergillus ochraceus Streptomyces rimosus Cunninghamella echinulata Beauveria bassiana Cunninghamella echinulata

C-4- and C-4′ Hydroxylations N-Monodemethylation, N-oxidation

Oxidative deamination to carboxylic acid

D-ring hydroxylations (unidentified position) N-Deisopropylation, oxidative deamination to alcohol or carboxylic acid, ring C-4 hydroxylation Reduction to sulfide and oxidation to sulfone N-Oxidation and N-demethylation

C-4 Hydroxylation N-Demethylation C-4 ring hydroxylation

Phencyclidine (hallucinogen)

Pheniramine Brompheniramine Chlorpheniramine (antihistamine) Primaquine (antimalarial)

Praziquantel (antihelminthic)

Propanolol (β blocker)

Sulindac (anti-inflammatory) Tamoxifen (anti-estrogen, antitumor)

Thymoxamine (α-adrenergic blocking agent)

Tranylcypromine (antidepressant)

Modified after [10]. a Additional reaction types that are reported only for the mammalian or only for the microbial systems investigated are not listed. For these data please refer to [10].

Nocardia corallina Gliocladium roseum Cunninghamella elegans Cunninghamella blakesleena Streptomyces rimosus Mortierella isabellina Mucor rouxii Cunninghamella echinulata

Cunninghamella echinulata Curvularia falcate Streptomyces griseus Beauveria bassiana Cunninghamella echinulata Cunninghamella elegans

ω-Methyl group oxidation Oxidative cleavage of the ketonic side-chain

Parbendazole (antihelminthic) Pentoxifyline (hemorheological agent)

9.6 Correlation of Microbial Reactions with Human CYP Isozyme-Specific Reactions 223

ATCC 10600 DSM 875 DSM 1906 DSM 1908 ATCC 38850 DSM 62752 ATCC 10002 DSM 905 ATCC 12938 DSM 43230 DSM 30121 ATCC 31560

Aspergillus alliaceus (F) Beauveria bassiana (F) Cunninghamella blakesleeana (F) Cunninghamella elegans (F) Curvularia lunata (F) Mortierella ramanniana (F) Penicillium chrysogenum (F) Rhizopus oryzae (F) Rhizopus stolonifer (F) Rhodococcus sp. (Nocardia corallina) (B) Serratia marcescens (B) Streptomyces violascens (B)

Phenacetine (CYP1A2) nd − ++ ++ + − + − − − − +

Caffeine (CYP1A2) − − − − − − − − − − − −

Theophylline (CYP1A2) nd nd − − nd nd nd nd nd − + nd

+ + ++ ++ nd + nd − nd − − +

Tolbutamide (CYP2C9)

Model drugs as CYP substrates (Test substrate for human CYP isozyme)

nd − + − ++ − − − − − − −

Desipramine (CYP2D6)

Out of the 25 evaluated microorganisms only those showing significant transformation of at least one model drug are reported. ++, rapid product formation; +, slow product formation; −, no conversion; nd, not determined. Pasel, S., Kittelmann, M., Ghisalba, O., Novartis unpublished results.

Strain (collection number)

Strain name F: Fungus B: Bacterium

− − + ++ nd − nd + nd − − ++

Nifedipine (CYP3A4)

Table 9.5 Bacterial and fungal cytochrome P450 reaction screening

− − − − − − − + + + + +

Chlorzoxazone (CYP2E1)

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9.7 Novartis Research Examples of Microbial Hydroxylations

spectrum: for the same microbes completely different behaviors may be found with respect to other known substrates of the same human CYP isozyme. In many cases such comparative studies cannot replace a proper microbial biocatalyst screening dedicated to the specific CYP substrate to be converted. An excellent and very instructive example of an in-depth comparative investigation of bacterial and human CYP reactions was recently published for the various types of testosterone hydroxylations [15]. In this work 213 bacterial cytochrome P450 genes were collected (about 70% from actinomycetes) and expressed based on an E. coli expression system [16] to test their hydroxylation activities with testosterone. Stereoselective monohydroxylations of testosterone were identified with 24 of those bacterial CYPs at the 2α-, 2β-, 6β-, 7β-, 11β-, 12β-, 15β-, 16α-, and 17-positions (17-hydroxylation yields 4-androstene-3,17-dione). However, the hydroxylation site usage of the bacterial CYPs was not the same as that of human CYPs. The 2α-, 2β-, 6β-, 11β-, 15β-, 16α-, and 17-hydroxylations are reactions common to both bacterial and human CYPs. Most of the testosterone hydroxylations catalyzed by bacterial CYPs are on the β face. The 1β-, 6α-, 7α-, 15α-, and 16β-hydroxylations (reported in human liver microsomes) are not found with the bacterial CYPs investigated. The most versatile bacterial CYP hydroxylating testosterone at positions 2β-, 6β-, 11β-, 12β-, 15β-, and 17 was reported to be CYP219A1 from Novosphingobium aromaticivorans DSM 12444. In comparison, CYP3A4, the most abundant P450 in human liver and small intestine, can hydroxylate testosterone in 10 different positions [17]. Microbial conversion of steroid precursors – with whole cells or ruptured cells – is widely used in large-scale synthesis of steroid drugs, mostly using fungal strains [18–21].


The Novartis Bioreactions Laboratories (formerly Ciba-Geigy Bioreactions Laboratories) have had positive experience with microbial hydroxylation of drug precursors, synthetic drugs, and biologically active natural products over a long period of time. Some typical and instructive examples are present here. 9.7.1 Preparation of 10,11-Epoxy-carbamazepine and 10,11-Dihydro-10-hydroxy-carbamazepine

Carbamazepine (CBZ) and 10,11-dihydro-10-oxo-CBZ are the active ingredients of the antiepileptic pharmaceuticals Tegretol and Trileptal. In humans, CBZ is metabolized mainly via 10,11-epoxy-CBZ and 10,11-dihydro-10-oxy-CBZ via 10,11dihydro-10-hydroxy-CBZ (10-OH-CBZ) (Fig. 9.3). Both metabolites also have strong anticonvulsive activities. For metabolic and pharmacokinetic studies these two CBZ-metabolites were required in sufficient quantities.

225

226


Fig. 9.3 Epoxidation of CBZ to 10,11-epoxy-CBZ and hydroxylation of H2-CBZ to 10-OH-CBZ.

Within a limited microbial screening (with whole cells and 1 g L −1 of CBZ) starting from 11 bacterial and 10 fungal strains (known from the literature to be “versatile metabolizers”) only two bacterial strains were identified that epoxidize CBZ to 10,11-epoxy-CBZ: Streptomyces diastatochromogenes ATCC 31561 and Streptomyces violascens ATCC 31560. For the scale-up of the reaction (0.5 g L −1 of CBZ in an MBR-Minireactor with 2 L working volume) the more active strain S. violascens was used and 265 mg of pure 10,11-epoxy-CBZ could be isolated [22]. In a parallel screening (with whole cells and 1 g L −1 of H2-CBZ) starting from 37 bacterial and 11 fungal strains in total 10 strains all belonging to the genus Streptomyces were identified that hydroxylate 10,11-dihydro-CBZ (H2-CBZ) to 10,11-dihyhdro-10-hydroxy-CBZ (10-OH-CBZ). The five most active strains were: Streptomyces violascens ATCC 31560, Streptomyces griseus ATCC 13273, Streptomyces griseus subsp. griseus DSM 40855 = ATCC 10137, Streptomyces griseus subsp. griseus DSM 40881 = ATCC 13963 and Streptomyces sp. DSM 40865 = ATCC 13279. For the scale-up of the reaction (0.5 g L −1 of CBZ in shake flasks with 4 × 100 mL medium) again the S. violascens was used and 104 mg of pure 10-OHCBZ could be isolated [22]. With S. violascens the culture conditions for the hydroxylation of H2-CBZ were improved by medium optimization. The best conversion yields were obtained at pH 6 and with soybean flour as a complex nutrient. In the best experiment a conversion yield of 78% and an average reaction rate of 210 mg L −1 day−1 were reached. Streptomyces violascens and Streptomyces griseus are described as having a cytochrome P450-dependent hydroxylating system with a broad substrate range [23, 24], inducible by soybean flour and genistein [23].


Fig. 9.4 Microbial hydroxylation of a highly water-soluble analog of H2-CBZ.

In a later stage of our investigation, the enantioselectivity of the microbial hydroxylation in position 10 was determined and it was found that the five Streptomyces strains show different degrees of selectivity but all preferentially produce the S-enantiomer. The following S/R-ratios were found for the individual strains by chiral analysis of the reaction product: Streptomyces sp. DSM 40865 (S/R = 92/8), Streptomyces griseus DSM 80855 (82/18), Streptomyces griseus DSM 40881 (82/18), Streptomyces griseus ATCC 13273 (80/20), and Streptomyces violascens ATCC 31560 (57/43). Based on these results, the further scale-up operations were performed with Streptomyces sp. DSM 40865. Twelve grams of 10-OH-CBZ were isolated from a 20 L biotransformation batch (MBR fermenter) (Kittelmann, M., Ghisalba, O. unpublished results). Attempts were made to further enlarge the scale of H2-CBZ hydroxylation. However such a scaling-up operation is hampered by the relatively low solubility of the substrate H2-CBZ and the product 10-OH-CBZ allowing only final product concentrations of 0.8 g L −1. In order to overcome this limitation, a highly watersoluble succinylated analog of H2-CBZ was synthesized (Fig. 9.4). In an additional limited screening program Streptomyces sp. DSM 40307 was found to quantitatively hydroxylate this highly soluble compound at a concentration of 10 g L −1 (Laumen, K., Kittelmann, M., Ghisalba, O., unpublished results). To obtain the hydroxylated H2-CBZ = 10-OH-CBZ from this product the solubilizing group would have to be substituted by a carbamoyl group. Such a route would, in principle, be feasible on a production scale, but was not, however, further developed by Ciba-Geigy/Novartis. 9.7.2 Preparation of 4-(4′-Hydroxyanilino)-5-anilinophthalimide and 4,5-Bis-(4′-hydroxyanilino)-phthalimide by Microbial Hydroxylation

Dianilinophthalimides (DAPHs) were identified as a novel class of potent and selective inhibitors of epidermal growth factor receptor protein tyrosine kinase (EGF-R PTK) with a high degree of selectivity compared with other tyrosine/ threonine kinases. This class therefore offers potential candidates for further optimization [25].

227

228


Fig. 9.5 Hydroxylation of DAHP to 4-(4′-hydroxyanilino)-5anilinophthalimide and 4,5-bis-(4′-hydroxyanilino)-phthalimide by microbial cells.

Monohydroxylated DAPH derivatives had to be made available in sufficient quantity for biological and pharmacokinetic studies. A microbial screening with collection strains from 30 different microbial genera and newly isolated strains (in total 153 strains) revealed that two fungal strains, Beauveria bassiana DSM 1344 = ATCC 7159 and Cunninghamella elegans DSM 1908 = ATCC 9245, as well as three newly isolated bacterial strains belonging to the genus Streptomyces/Actinomyces were able to hydroxylate 4,5dianilinophthalimide (DAPH, CGP52411) (250 mg L −1) to 4-(4′-hydroxyanilino)-5anilinophthalimide. Cunninghamella elegans DSM 1908 was found to be the most active of those biocatalysts and was also able to form the dihydroxy derivative, 4,5-bis-(4′-hydroxyanilino)-phthalimide (Fig. 9.5) [26]. After optimization of the medium and of the biotransformation conditions the reaction for the monohydroxylated biotransformation product was carried out on preparative scale with Cunninghamella elegans (0.25 g L −1 of DAPH in an MBRMinireactor with 2 L working volume) and 227 mg of pure 4-(4′-hydroxyanilino)5-anilinophthalimide could be isolated. Two 20 L fermentations were subsequently performed to produce 5 g of this compound. The dihydroxylated product 4,5-bis(4′-hydroxyanilino)-phthalimide can be produced with the same strain in a modified medium up to a concentration of 50 mg L −1 [26].

9.8 Microbial Oxidation of Natural Products

Typical oxidative “metabolic” reactions in the context of modification of natural products are: Baeyer–Villiger oxidation, oxidation of aromatic and non-aromatic centers (i.e. hydroxylations), oxidoreductions and O- and N-dealkylations. Three

9.9 Conclusions

microbial pathways for N-dealkylation are known: via oxidases, dehydrogenases, or CYPs [27]. Substrates (or groups) for microbial Baeyer–Villiger oxidations are steroids, alkanes, and cyclic ketones. Examples include the side-chain cleavage of 20-ketosteroids by Fusarium sp., the conversion of testosterone/progesterone to testolactone by Aspergillus sp. or Penicillium sp. and the conversion of 4-hydroxyandrost-4-ene-3,17-dione to the corresponding D-ring lactone by Cylindrocarpon radicicola. Baeyer–Villiger monooxygenases (BVMOs) are flavoenzymes that catalyze a wide variety of oxidative reactions such as stereo- and enantioselective Baeyer–Villiger oxidations and sulfoxidations. They are found in biosynthetic pathways in many different organisms and also in microbial degradation pathways [27, 28]. 9.8.1 Microbial Hydroxylation and Epoxidation of Milbemycins

Many examples of microbial hydroxylations of natural products can be found in the literature. A typical example from our own laboratory is the diastereoselective hydroxylation of milbemycin derivatives showing high antiparasitic activity [29]. The task was to hydroxylate milbemycin A4 at the 13β-position. This derivative was requested as a starting material for chemical derivatization (i.e. the synthesis of 13β-alkoxy, alkylthio, acyloxy, carbamoyloxy and alkyl milbemycin derivatives). In a very limited screening with three bacterial and eight fungal strains known to have potential hydroxylation activity, Streptomyces violascens ATCC 31560 was identified as a suitable biocatalyst performing the requested 13β-hydroxylation of milbemycin A4 (Fig. 9.6). 14,15-Epoxymilbemycin A4 was formed via a parallel reaction as a byproduct. The conversion and the product ratio of 13β-hydroxy vs. 14,15-epoxymilbemycin A4 could be improved by the addition of organic solvents. The best results (with 2 g L −1 milbemycin A4 ) (i.e. 91% overall conversion with a product ratio of 92% 13β-hydroxymilbemycin A4 and 8% 14,15- epoxymilbemycin A4 ) were obtained by adding 2.5% DMSO to the culture broth. In analogous biotransformations with the same microorganism and milbemycins A3 (methyl instead of ethyl in position 22] and D (isopropyl in position 22), the resulting main products were the corresponding 14,15-epoxides and the corresponding 13βmilbemycins were present as byproducts. The 13β-hydroxy-/14,15-epoxy-product ratios under identical conditions were found to be 70:30 for milbemycin A4, 40:60 for milbemycin A3 and 29:71 for milbemycin D.

9.9 Conclusions

Microbial biocatalysts are a very useful complement to the mammalian/human metabolic enzymes performing phase I oxidative metabolism, especially when

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230


Fig. 9.6 Microbial hydroxylation and epoxidation of milbemycins.

the synthesis of larger amounts of metabolites (i.e. above gram-scale) is requested. However, unfortunately there are also some restrictions to the microbial “alternative” because microbial (bacterial, fungal) oxidative metabolism is not always directly comparable to mammalian systems. Many types of mammalian metabolites can be successfully generated with microbial systems. It must be borne in mind, though, that in a significant number of cases specific mammalian metabolites cannot be generated with microbial systems and in other cases the microbes produce additional types of metabolites that are not found with the mammalian systems. In the worst case for a given drug candidate (substrate) the metabolic patterns generated by mammalian/human DME systems can be totally different from the microbial ones. Given the present state of knowledge, predictions of oxidative reactions with microbes in comparison to mammalian systems can only be tentative, so every biotransformation task designed to produce a required metabolite must be regarded as an individual case and laborious screening carried out. Practical limitations in the biotransformation process may include the low turnover numbers of the involved oxygenases (Note: A compilation of selected known k kat values can

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10 Azerad, R. Microbial models for drug metabolism. Adv Biochem Eng/Biotechnol 1999, 63, 169–218. 11 Srisilam, K., Veeresham, C. Biotransformation of drugs by microbial cultures for predicting mammalian drug metabolism. Biotechnol Adv 2003, 21, 3–39. 12 Smith, R.V., Rosazza, J.P. Microbial systems for study of the biotransformations of drugs. Biotechnol Bioeng 1975, 17, 785–814. 13 Smith, R.V., Rosazza, J.P. Microbial models of mammalian metabolism. J Pharm Sci 1975, 64, 1737–1759. 14 Smith, R.V., Acosta, D. Jr., Rosazza, J.P. Cellular and microbial models in the investigation of mammalian metabolism of xenobiotics. Adv Biochem Eng 1977, 5, 70–100. 15 Agematu, H., Matsumoto, N., Fujii, Y., Kabumoto, H., Doi, S., Machida, K., Ishikawa, J., Arisawa, A. Hydroxylation of testosterone by bacterial cytochromes P450 using the Escherichia coli expression system. Biosci Biotechnol Biochem 2000, 70, 307–311. 16 Arisawa, A., Kumeda, A. World Patent WO2003/087381 (October 23), 2003. 17 Evans, W.E., Relling, M.V. Pharmacogenomics: translating function genomics into rational therapeutics. Science 1999, 286, 487–491. 18 Mahato, S.B., Mukherjee, A. Microbial transformation of testosterone by Aspergillus fumigatus. J Steroid Biochem 1984, 21, 341–342. 19 Smith, K.E., Latif, S., Kirk, D.N. Microbial transformation of steroids – II. Transformations of progesterone, testosterone and androstenedione by Phycomyces blakesleanus. J Steroid Biochem 1989, 32, 445–451. 20 Smith, K.E., Latif, S., Kirk, D.N. Microbial transformation of steroids – IV. Transformations of testosterone and

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25

androsterone by Botryosphaerica obtusa. J Steroid Biochem 1990, 35, 115–120. Kolek, T., Swizdor, A. Biotransformations XLV: Transformations of 4-ene-3-oxo steroids in Fusarium culmorum culture. J Steroid Biochem Mol Biol 1998, 67, 63–69. Kittelmann, M., Lattmann, R., Ghisalba, O. Preparation of 10,11-epoxycarbamazepine and 10,11-dihydro-10hydroxy-carbamazepine by microbial epoxidation and hydroxylation. Biosci Biotechnol Biochem 1993, 57, 1589–1590. Sariaslani, F.S., Kunz, D.A. Induction of cytochrome P450 in Streptomyces griseus by soybean flour. Biochem Biophys Res Commun 1986, 141, 405–410. Trower, M.K. Sariaslani, F.S., Kitson, F.G. Xenobiotic oxidation by cytochrome P-450-enriched extracts of Streptomyces griseus. Biochem Biophys Res Commun 1988, 157, 1417–1422. Trinks, U., Buchdunger, E., Furet, P., Kump. W., Mett, H., Meyer, T., Müller, M., Regenass, U., Rihs, G., Lydon, N.B., Traxler, P. Dianilinophthalimides: Potent and selective ATP-competitive inhibitors of the EGF-receptor protein

26

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tyrosine kinase. J Med Chem 1994, 37, 1015–1027. Weidner, S., Goeke. K., Trinks, U., Traxler, P., Ucci-Stoll, K., Ghisalba, O. Preparation of 4-(4′-hydroxyanilino)-5anilinophthalimide and 4,5-bis-(4′hydroxyanilino)-phthalimide by microbial hydroxylation. Biosci Biotechnol Biochem 1999, 63, 1497–1500. Sima Sariaslani, F., Rosazza, J.P.N. Biocatalysis in natural product chemistry. Enzyme Microb Technol 1984, 6, 242–253. Kamerbeek, N.M., Janssen, D.B., van Berkel, W.J.H., Fraaije, M.W. Baeyer-Villiger monooxygenases, an emerging family of flavin-dependent biocatalysts. Adv Synth Catal 2003, 345, 667–678. Ramos Tombo, G.M., Ghisalba, O., Schär, H.P., Frei, B., Maienfisch, P., O’Sullivan, A.C. Diastereoselective microbial hydroxylation of milbemycin derivatives. Agric Biol Chem 1989, 53, 1531–1535. Duetz, W.A., van Beilen, J.B., Witholt, B. Using proteins in their natural environment: potential and limitations of microbial whole-cell hydroxylations in applied biocatalysis. Curr Opin Biotechnol 2001, 12, 419–425.

Additional Review References for Further Reading Azerad, R. Microbiological hydroxylations: Myths and realities. Chimia 1993, 47, 93–99. Guengerich, P.F. Common and uncommon cytochrome P450 reactions related to metabolism and chemical toxicity. Chem Res Toxicol 2001, 14, 611–650. Guengerich, P.F. Cytochrome P450 enzymes in the generation of commercial products. Nat Rev Drug Discov 2002, 1, 359–366.

Mansuy, D. Cytochromes P-450 and model systems: great diversity of catalysed reactions. Pure Appl Chem 1994, 66, 737–744. Mansuy, D. The great diversity of reactions catalysed by cytochromes P450. Comp Biochem Physiol C 1998, 121, 5–14. Wrighton, S.A., Stevens, J.C. The human hepatic cytochromes P450 involved in drug metabolism. Crit Rev Toxicol 1992, 22, 1–21.

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10 Recombinant Yeast and Bacteria that Express Human P450s: Bioreactors for Drug Discovery, Development, and Biotechnology Steven P. Hanlon, Thomas Friedberg, C. Roland Wolf, Oreste Ghisalba, and Matthias Kittelmann

Summary

A wide variety of important pharmacological and toxicological properties of xenobiotics are determined by cytochrome P450-mediated metabolism. Prediction of these pathways in humans and of the P450 isoenzymes involved constitutes an essential part of drug development. Recently a variety of recombinant expression systems have been employed to predict the P450-mediated metabolism of drug candidates in humans. These include insect, bacterial, yeast, and mammalian models which all have been successfully employed recently on a small scale in drug development. However, this article focuses mainly on the comparison of bacterial and yeast systems for semi-preparative biooxidations. We demonstrated that the catalytic properties of several P450 isoforms when expressed in either Saccharomyces cerevisiae or Escherichia coli were rather similar and faithfully mimicked metabolic pathways of xeno- and endobiotics in human liver. However, taking technical considerations into account, the high level of functional expression of P450s achieved in bacteria make this system, which is patented, ideally suited for large-scale pharmaceutical biooxidations. These include the semipreparative generation of metabolites for structural determinations and, importantly, also for drug safety studies. This suitability is evidenced by the industrial applications presented in this review. These were conducted by two pharmaceutical companies, involving structurally diverse compounds, and utilized different bioreactor techniques. The strategies employed resulted in highly efficient, semi-preparative biooxidations of drugs and drug candidates, showing the great potential of these systems to improve and facilitate drug discovery and development.

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10 Recombinant Yeast and Bacteria that Express Human P450s

10.1 Background 10.1.1 Importance of Recombinant P450s for Drug Development

The drug discovery and development process is scientifically complex, incorporating many research areas, and is inherently full of challenges and risk. It is therefore expensive and time consuming, taking typically up to 15 years and consuming up to 500 million euro per approved drug. The entire process can be divided into drug discovery and preclinical drug development, followed by clinical development and approval. Typically, new chemical entities (NCEs) are promoted from discovery into preclinical development and if they succeed to pass all hurdles, they are then submitted for investigational drug (IND) applications and eventually enter the various phases of clinical development. The attrition (failure) rate is very high in the preclinical stages, which are designed to predict the therapeutic properties (desired and adverse) in humans. An important part of preclinical development is the prediction of drug metabolism and its implications for drug safety and pharmacokinetics in humans. The metabolism of a drug proceeds in two phases with the first phase being of greatest importance for the metabolism of pharmaceuticals. This step is mainly catalyzed by a large family of highly versatile cytochrome P450 monooxygenases (CYPs) which differ in their substrate-, regio-, and stereoselectivity. To ensure the maximum therapeutic value and safety of pharmaceuticals in humans, the metabolism of a NCE is first tested in animals or in vitro systems that mainly rely on animal tissues [1, 2]. Besides being often considered ethically and economically problematic, animal-based systems have limitations partially due to large species differences in the P450s involved in drug metabolism [3, 4]. It is therefore essential to develop systems that are based on material derived from human tissues. These should significantly shorten the period between the discovery of drugs and their introduction into the market and simultaneously allow more accurate extrapolations to humans. Models for human drug metabolism can roughly be classified into those that try to mimic the entire cascade of drug-metabolizing events, and those that imitate only a limited set of reactions. The former models (complex systems) include human hepatocyte and hepatoma cell cultures. The latter (simple systems) include heterologous in vivo and in vitro expression systems as well as enzymes purified from tissues. A disadvantage of the complex models is that drug metabolism is in most cases altered as compared to liver, either due to culture conditions, or in the case of hepatoma cells, due to phenotypic transformation [5, 6]. In addition, these models are only poorly suited for large-scale biooxidation reactions. Simple models, employing microorganisms such as bacteria or yeast, are useful for characterizing specific steps in the metabolism of drugs and also the enzymes involved in them. These systems also yield recombinant enzymes for antibody production [7], which allows quantitation of drug-metabolizing enzymes in

10.1 Background

human tissues, including antibody inhibition studies. Combined with the knowledge of the role of a particular enzyme in drug metabolism, as derived from the use of bacterial or yeast expression systems, this information can be used to predict the metabolic pathways of drugs in vivo. E. coli expression systems for P450 have been employed with great success to predict polymorphic drug oxidations [8–11]. Furthermore, these bacterial models provide a powerful means to predict drug/drug interactions which are frequently observed in vivo [12] and sometimes result in life-threatening adverse drug reactions. Bacterial expression systems for P450s are also highly suitable to express P450s mutated by site-directed mutagenesis to delineate amino acid residues in the catalytic center of these enzymes that are important for their catalytic activity [13, 14]. This information, combined with the recently available crystal structure of P450s, which was derived from bacterially expressed human P450s [15], is necessary to establish reliable in silico models of these proteins that will ultimately revolutionalize in silico predictions of drug metabolism [16]. Last but not least, microbial expression systems for mammalian P450s are ideally suited to the generation of drug metabolites on a semi-preparative scale as evidenced in this review and from the literature [17, 18]. This strategy will greatly facilitate the structural elucidation of the metabolites and drug safety testing [19] employing the resulting metabolites. 10.1.2 Fundamentals of Heterologous Expression in Bacteria

Escherichia coli has been most frequently used for the bacterial expression of human drug-metabolizing enzymes. E. coli is an attractive system because high levels of expression as well as growth to very high cell densities can be achieved [20]. In addition, E. coli is easily manipulated and a wide variety of strain variants and vectors with powerful promoters are available. A limitation of the bacterial systems is that in almost all cases mammalian cDNAs have to be modified before they can be expressed [20]. Aside from trimming the 5′ and 3′ untranslated regions of the P450 cDNAs, the region around the initiation codon of protein biosynthesis should be modified to remove the rigid secondary structures that can occur in mammalian mRNAs. We have developed a strategy that circumvents changes in the coding region of the P450s by inframe fusions of a modified ompA leader sequence to the various human P450 cDNAs. This leader is removed during P450 synthesis, thus releasing the native P450. To obtain a functional monooxygenase complex in E. coli, it is also necessary to coexpress cytochrome P450 reductase, which is lacking in this organism and which supplies electrons to P450s. Several strategies have been developed to this end. Because our strategies and a detailed methodology for the bacterial expression of human P450s on a small scale has been given in various reviews and original articles [11, 17, 21–24] and is patented (US 6,566,108; WO98/02554), it will only be touched upon here. We will focus more on the utilization of these microorganisms for biooxidation on a larger scale.

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10.1.3 Fundamentals of Heterologous Expression in Yeast

Bakers yeast, Saccharomyces cerevisiae, has been almost exclusively used for the heterologous expression of drug-metabolising enzymes in yeast. S. cerevisiae has proved suitable for the production of a wide variety of eukaryotic proteins in both basic research and industrial and pharmaceutical application [25]. This unicellular organism has some of the advantages of the bacterial expression systems but the additional advantage of being a eukaryotic cell with many similarities to mammalian cells in protein synthesis and processing and membrane compartmentalization. Two types of yeast vector are available [26]: episomally replicating vectors and vectors with the potential for genomic integration. Most episomal vectors contain sequences from the 2 μm endogenous yeast plasmid that confer high copy number and stable maintenance. Integrating plasmids lack a yeast origin of replication but contain regions of homology with the yeast genome flanking the cDNA cloning site. The stable integration of an ancillary protein, such as cytochrome P450 reductase, required for functional cytochrome P450 expression [27], increases the versatility of yeast for the subsequent expression of further proteins. S. cerevisiaebased expression biotechnology benefits from the availability of numerous selection markers for gene transfer, making it possible to cotransfect several cDNAs. Interestingly, expression of the steroidogenic CYP17A in the yeast Yarrowia lipolytica yielded an active P450 enzyme in the absence of coexpressed recombinant reductase [28]. This seems to be a feature of the CYP17A isoform, which also does not require exogenous reductase in E. coli, as another human P450 isoform (CYP1A1) expressed in Yarrowia required coexpression of P450 reductase to be catalytically active [29]. Similarly, the presence of mammalian P450 reductase was required for the catalytic activity of CYP2D6 when expressed in Pichia pastoris [30]. There are also several powerful regulatable S. cerevisiae promoters that can yield high levels of heterologous protein expression. However, some features of mammalian cDNAs are not optimal for the expression of proteins in S. cerevisiae. It has been shown that efficient expression of several proteins, including cytochrome P450s, requires the deletion of most of the 5′ untranslated region [27]. For example, a CYP1A1 cDNA under the control of the GAL10-CYC1 promoter containing either 15 bp or 5 bp of 5′ untranslated region, yielded 1 μg and 6 μg of functional CYP1A1 per mg of microsomal protein respectively [31]. Even though it does not appear to be necessary to change the coding region of mammalian cDNAs for their expression in S. cerevisiae, certain mammalian proteins are difficult to express unless the amino acid sequence is modified. An example is the rat cytochrome P450 reductase, which could not be stably expressed unaltered in S. cerevisiae. However, upon fusion of the N-terminal sequence of this protein with the N-terminus of the P450 reductase from S. cerevisiae, high levels of the rat P450 reductase were achieved [32]. Coexpression of mammalian P450 reduc-

10.2 Comparison of P450 Levels and Enzymic Activities in Various Models

tase and P450s in S. cerevisiae is required to achieve a catalytically highly active monooxygenase system, since the host P450 reductase couples poorly with mammalian P450s [33]. We have established a functional monooxygenase system in S. cerevisiae by stable integration of the P450 reductase linked to the 3′ phosphoglycerol kinase (PGK) promoter into the yeast genome. The resulting strain had a cytochrome c reductase activity of more than 200 nmol min−1 mg−1 microsomal protein which was 20-fold higher than the activity in the parental strain. The recombinant strain was subsequently transformed with an episomally replicating vector containing the various P450 cDNAs under the control of the powerful constitutive PGK promoter. In the absence of human reductase, the human P450s were catalytically inactive, however upon expression of this ancillary factor, CYP3A4 and CYP2D6 in S. cerevisiae displayed an activity towards prototypical substrates such as testosterone and bufuralol respectively. Interestingly the expression level of the various P450 isozymes varied widely, with the level of CYP2C9 being only 91 pmol mg−1 microsomal protein whereas the level of CYP3A4 in microsomes was found to be 500 pmol mg−1 (Table 10.1).


Table 10.1 lists the P450 isozymes that have been expressed by us in the different microorganisms, their expression levels, and their enzymatic characterization. P450 isozymes that have been expressed in at least two systems (bacteria and yeast) are listed in bold. The levels of recombinant P450s (pmol min−1 mg−1 protein) in the different cellular systems were surprisingly similar. However, we found that the level of some P450s in E. coli can be increased to about 1000 pmol mg−1 protein when growing the cells in the controlled environment of a biofermenter. This level represents approximately 5% of the bacterial membrane protein, which translates into a yield of 1000 nmol P450 L −1 of culture. Similarly it has been shown by others that the yield of P450s in S. cerevisiae can be improved to about 500 nmol L −1 of culture when using vectors that are stably maintained in rich medium, which allows the cells to grow to very high cell densities [33]. Therefore E. coli and S. cerevisiae are ideally suited for the production of sufficient quantities of enzymes for structural studies. The testosterone 6β-hydroxylase and the bufuralol 1′-hydroxylase activity (pmol min−1 mg−1) found in membranes isolated from bacteria which expressed CYP3A4 or CYP2D6 respectively were 2- and 30-fold respectively higher than in human liver microsomes (see Table 10.1 footnote). Importantly, these activities in bacterial membranes were 120- and 6-fold higher than the activities found in yeast membranes (Table 10.1). This also strongly suggests that, apart from the ease of handing, the bacterial systems are far superior to the yeast-based systems for large-scale biooxidations.

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Table 10.1 Catalytic activities and levels of cytochrome P450s expressed in E. coli and S. cerevisiae.

Expression system

Expressed P450 isoform

P450 expression level[a] (pmol mg-1)

Yield (nmol L-1)

Catalytic activity[b] (pmol min -1 mg-1)

E. coli

1A1 1A2 1B1 2A6 2B6 2C8 2C9 2C19 2D6 2E1 3A4 3A5 4A11 2C9 2D6 3A4

235 204 195 190 25 1700 286 n.a. 210 70 300 150 80 91 100 500

170 218 287 150 30 440 498 40 370 190 210 250 70 0.9 2.6 4.4

725 63 180 550 37 n.d. 2 400 n.a. 1 200 1 360 12 000 700 1 600 n.d. 209 104

S. cerevisiae

Turnover number (min -1)

3.6 0.3 0.92 2.9 1.2 n.d. 8.7 1.43 5.7 30 40 4 20 n.d. 8.6 1.5

Km (mmol L-1)

n.a. n.a. n.a. n.a. n.a n.a. n.a. n.a. 11.1 n.a. 69 n.a n.a. n.d. 11.1 99

Heterologous expression levels for P450s were determined in whole bacteria or yeast cells (yield) or in bacterial membranes or yeast microsomes (pmol mg−1 protein) and represent typical values. Enzyme activities were determined in bacterial membranes or yeast microsomes. P450 isoforms that were expressed in E. coli as well as in yeast are in bold. The various P450 isozymes were coexpressed with human P450 reductase. The cytochrome c reductase activities found in E. coli strains coexpressing P450s and P450 reductase (hOR) were typically 400–1200 nmol cytochrome c reduced min−1 mg−1 membrane protein with the exception of strains expressing CYP1A2 for which this value was 70–120. In S. cerevisiae coexpressing P450s and hOR the levels of reductase were 200–250 nmol cytochrome c reduced min−1 mg−1 membrane protein. n.a., not available. a The following enzyme activities have been determined for the different P450 isoenzymes (the typical specific activities reported in literature for human hepatic microsomes are given in brackets): CYP1A1 and CYP1A2, 7-ethoxyresorufi n O-dealkylase (95 pmol min−1 mg−1); CYP2B6, diazepam-demethylase; CYP2C9, diclofenac 4′-hydroxylase; CYP2D6, bufuralol 1′- hydroxylase (70 pmol min−1 mg−1); CYP2E1, chlorzoxazone 6-hydroxylase (1500–4000 pmol min−1 mg−1); CYP3A4 and CYP3A5, testosterone 6 ß-hydroxylase (5000 pmol min−1 mg−1); CYP4A11, lauric acid omega-hydroxylation; P450 reductase, cytochrome c (230 nmol min−1 mg−1). b The P450 content was determined spectrophotometrically and is expressed as pmol mg−1 membrane protein.

For pharmacokinetic investigations it is essential that the catalytic properties of the recombinant P450 isozymes are similar across the various recombinant models. The data presented in Table 10.1 show that the K m values of the CYP3A4mediated 6β-hydroxylation of testosterone and the CYP2D6-mediated 1′hydroxylation of bufuralol were closely similar in the two models developed in this work. These values are also in good agreement with the K m reported for CYP2D6 expressed either in the baculovirus system or in human lymphoblastoid


cells which were 4.7 μmol L −1 and 18 μmol L −1 respectively [34, 35]. Similarly the K m of CYP3A4 expressed in the two models presented here agrees with the K m reported for CYP3A4 expressed in a baculovirus system, which was 56 μmol L −1 [36]. In many cases, however, the catalytic properties of recombinant P450s cannot be directly compared with those of P450s in human liver microsomes, since here several P450s can contribute to the metabolism of a given substrate. For example, it is known that the bufuralol 1′-hydroxylation is catalyzed by CYP2D6 but also with a lower affinity by CYP1A2 [35]. Similarly, the testosterone 6βhydroxylation is catalyzed not only by CYP3A4 but also by CYP3A5 [36]. It can be said that except for CYP1A2, the substrate turnover numbers (min−1) of the P450 isozymes expressed in E. coli differed by a factor of less than four from those found in liver (Table 10.1 footnote). The relatively low turnover number of bacterially expressed CYP1A2 for the O-dealkylation of 7-ethoxyresorufin is most likely due to the low level of P450 reductase in this particular strain of E. coli, which was at least 4-fold lower than in the other E. coli lines. Lower levels of reductase may have been also partially responsible for the low substrate turnover number of CYP3A4 expressed in S. cerevisiae. Another reason for the low enzyme activity of CYP3A4 in yeast membranes could be the absence of cytochrome b5, which has been shown to stimulate the activity of CYP3A4 towards several substrates. It should be noted that another group has coexpressed CYP3A4 together with cytochrome b5 and P450 reductase in S. cerevisiae [37], however the turnover number given in that report is similar to our value for the testosterone 6β-hydroxylase activity of CYP3A4 in the absence of cytochrome b5. Interestingly this ancillary factor is absent in E. coli in which CYP3A4 displayed a surprisingly high catalytic activity. One may speculate that an E. coli protein substituted for cytochrome b5 in stimulating the activity of CYP3A4 [21] and indeed simultaneous expression of human cytochrome b5 together with CYP3A4 and P450 reductase stimulated the testosterone 6β-hydroxylase activity of this P450 isoform only 1.6-fold compared with the activity in the absence of cytochrome b5 [38]. The substrate turnover numbers of the recombinant CYP3A4 and of the recombinant CYP2D6 expressed in E. coli appears to be rather similar to the substrate turnover number estimated for these enzymes in human liver microsomes, which are 55 min−1 [36] and 10 min−1 [39], respectively. However, one caveat in this comparison is that more than one P450 may metabolize a given substrate and that P450 isozymes are difficult to quantitate exactly in human liver microsomes. Since the catalytic properties (K m, turnover number) of the P450 isozymes are rather independent of the cellular system used for their expression, recombinant models could be very valuable in predicting P450-mediated metabolism in humans. However it remains to be seen if the substrate specificities of all the P450 isozymes are preserved in the expression systems. In order to address this our 14 industrial partners have assayed various recombinant P450s using a panel of isozyme-specific substrates [40, 41] and their and our data indicate that the catalytic properties (substrate-, regioselectivity and K m values) of the recombinant P450s are the same as their microsomal counterparts.

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10.3 Use of E. coli P450 Expression Systems in Bioreactors 10.3.1 General Considerations

Drug development requires the detailed characterization of metabolic pathways and their relevance for drug safety. This type of analysis necessitates that milligram quantities of metabolites need to become available, a task which until now had to be performed by elaborate chemical syntheses which become even more demanding when the metabolites result from stereoselective oxidations. It is obvious that recombinant P450s are ideally suited for the production of sufficient quantities of P450 metabolites, provided that the cellular system expresses these enzymes at high levels in a catalytically highly active form. In addition, it is important that the catalysis proceeds for sufficient time and that the fermentation system is suitable for scale-up. Our results show that the activity (expressed as pmol min−1 mg−1 membrane protein) of either CYP2D6 or of CYP3A4 in membranes isolated from recombinant bacteria were by a factor of 6- and 110-fold respectively higher than that of membranes isolated from yeast. In addition the yield of P450 in E. coli was between 50- and 100-fold higher than that in S. cerevisiae. Thus at least in our hands, E. coli appears to be more suitable as a bioreactor system than S. cerevisiae, even though it may be likely that under growth conditions which give higher yields of P450 [33] the yeast system could also be employed as biofermenter. Below we describe in detail several examples of the use of the bacterial expression systems that express human P450s in the production of drug metabolites in industrial settings. Note that the bioreactor conditions employed by the two companies were rather different and yielded the desired biooxidations of structurally highly diverse compounds at the desired efficiencies. 10.3.2 The Roche Experience 10.3.2.1 Background and Utility of P450 Systems in Pharma Research Roche obtained 14 recombinant human CYP450 isozymes coexpressed with NADPH-P450 reductase (CPR) in E. coli as part of a collaboration with the University of Dundee, Scotland (LINK Program). During the last 10 years these enzymes have been used extensively within Roche both as biocatalysts for the preparation of metabolites of drug candidates and for DMPK applications such as high-throughput CYP450 inhibition screening. The availability of state of the art fermentation facilities at our site has allowed the production of sufficient biomass for metabolite preparation on the milligram to gram scale for applications such as structure identification, analytical reference material, elucidation of toxicological properties, and so on. Equally important is the in-house expertise in downstream processing, as purification of the metabolites from biobroths,

10.3 Use of E. coli P450 Expression Systems in Bioreactors

although often neglected as a factor in biocatalysis, is often the most timeconsuming and difficult step in the process. The procedure for the preparation of one such metabolite in 100 mg amounts is described below. 10.3.2.2 Fermentation of Recombinant E. coli A generic fermentation protocol was developed which can be applied with minor modifications to all 14 recombinant CYP450s received from the LINK program. Fermentations were carried out at 30 ºC using modified Terrific Broth medium in 150 L Braun fermenters with a working volume of 100 L. Stirring speed and aeration was set such that dissolved oxygen (DO) decreased to below detectable levels after 4–6 h cultivation. Such low DO levels were also found to be necessary for formation of active CYP450 by other authors [42]. The inducer isopropyl β- D 1-thiogalactopyranoside (IPTG) was added when OD600nm reached 0.6–0.8, generally after 4 h growth. After 24–48 h fermentation time, depending on the CYP450 isoform, cells were harvested by continuous flow centrifugation, resuspended in a high sucrose-containing buffer and after shock freezing in dry ice were stored at −80 ºC. 10.3.2.3 Biotransformations Catalyzed by Recombinant CYP450 In an early protocol, the frozen E. coli biomass was thawed and the cells disrupted by treatment with lysozyme/EDTA followed by resuspension in hypotonic buffer. Biotransformations using this crude membrane preparation were carried out with an NADPH-regenerating system based on glucose-6-phosphate dehydrogenase. This system is rather costly in terms of reagents, and so we looked for a method in which the lysozyme treatment could be omitted and the cells used intact (i.e. still with capacity to regenerate NADPH). It was found that frozen cells which were thawed and resuspended in phosphate buffer pH 7.4 supplemented with a small amount of NADP had the capability to generate metabolites albeit with lower yields than when crude membranes were used. It was decided to optimize this process using CYP3A4 as the model enzyme as this isoform makes the largest contribution to drug metabolism in the liver [43] and is therefore the most important for metabolite synthesis. It was found that supplementation of the reaction buffer with a carbon source such as citrate or isocitrate greatly stimulated both hydroxylation (M1) and N-demethylation (M2) of an endothelin receptor antagonist (Table 10.2). Although the effect of addition of these Csources is substrate dependent, in most cases citrate and isocitrate are superior to glucose. A second example is the hydroxylation of chlozoxazone also by recombinant CYP3A4 shown in Fig. 10.1. In this case the yield of 6-OH chlorzoxazone was 8-fold higher with isocitrate than glucose and it can be postulated that NADPH regeneration is more efficient, probably due to the requirement for only one functioning enzyme, isocitrate dehydrogenase. It is possible that the freeze–thaw undergone by these cells adversely affects glycolytic/TCA cycle enzyme(s) and this may result in less efficient NADPH regeneration when glucose is used as cosubstrate.

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10 Recombinant Yeast and Bacteria that Express Human P450s Table 10.2 Effect of carbon source addition on

biotransformation of an endothelin receptor antagonist by frozen and thawed E. coli JM109 containing CYP3A4, P450 reductase and cytochrome b5. Carbon source

Metabolite M1 (mg L-1)

Metabolite M2 (mg L-1)

None Glucose Isocitrate Citrate

0 0 1.3 1.5

0 0.7 2.7 2.7

Reactions were carried out for 24 h at 27 ºC with shaking at 220 rpm in 24 deep-well plates containing in a total volume of 1.0 mL: 0.1 mol L −1 phosphate buffer pH 7, 1.2 nmol CYP3A4, 50 nmol NADP, and 50 nmol substrate.

Fig. 10.1 Hydroxylation of chlorzoxazone by “frozen and thawed” E. coli cells containing CYP3A4, NPR, and cytochrome b5, with addition of 20 mmol L−1 glucose or isocitrate. Reactions were carried out at 27 ºC with shaking at 220 rpm in 100 mL baffled flasks containing 10 mL 0.1 mol L−1 phosphate buffer pH 7.4: 8 nmol CYP3A4, 2.0 mmol chlorzoxazone, 0.5 mmol of either disodium citrate or glucose and 0.5 mmol NADP.

10.3.2.4 Preparation of N-Desethyl Amodiaquine N-Desethyl amodiaquine was required in multi-milligram amounts as an analytical reference for a CYP2C8 assay. CYP2C8, containing a modification at the Nterminus in which the first eight residues are replaced by those from bovine 17α-hydroxylase [44], was coexpressed with P450 reductase in E. coli JM109 using


a two plasmid system identical to that described for CYP2D6 [23]. Using the fermentation conditions described earlier, expression levels of CYP2C8 were typically 700–1000 nmol L −1 of culture after 24 h growth as determined by CO + red − red difference spectra. Cytoplasmic membranes prepared from such cells contained 1000–1200 pmol mg−1 CYP2C8 and CPR activity of 300–400 nmol cyt c red min−1 mg−1 protein. Optimization of amodiaquine biotransformation was carried out on the 10 mL scale in 100 mL shake flasks. Frozen E. coli cells were thawed slowly at room temperature and diluted in phosphate buffer pH 7.4 to a CYP2C8 concentration of 0.8 nmol mL −1. The concentrations of biocatalyst, substrate, citrate, and NADP were optimized using GALOP experimental design software (Forschungszentrum Jülich GmbH, Germany). Two rounds of optimization resulted in a doubling of substrate concentration from 0.2 to 0.4 mmol L −1 and a 4-fold reduction in the amount of biocatalyst (800 to 200 nmol L −1) required for complete conversion. Using the optimized conditions a 2 L reaction with 372 mg of amodiaquine dihydrochloride was carried out in 5 × 2 L flasks each containing 80 nmol CYP2C8. The biotransformation yield reached 93% after 32 h incubation as determined by HPLC (Fig. 10.2). The metabolite was purified by extraction followed by flash chromatography on silica gel. After concentration of the pure

Fig. 10.2 Biotransformation of amodaiquine by “frozen and thawed” E. coli cells containing CYP2C8 and NPR. Reactions were carried out at 27 ºC with shaking at 220 rpm in 2000 mL baffled flasks containing 400 mL 0.1 mol L−1 phosphate buffer pH 7.4: 80 nmol CYP3A4, 4.0 mmol amodiaquine, 20 mmol disodium citrate and 0.5 mmol NADP.

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fractions and drying of the residue, 172 mg of desethyl amodiaquine dihydrochloride was obtained with >99% purity in 55% overall yield. 10.3.3 The Novartis Experience 10.3.3.1 Introduction Already in the discovery phase, an understanding of the metabolic pathways of the lead drug candidates can be highly useful for improving the overall pharmacokinetic profi les of compounds in a lead structural series [45]. Although HPLCMS provides a powerful tool for the identification of low concentrated metabolites in solution, it is often still necessary to synthesize and isolate the metabolites on milligram to hundred milligram scale for final structure confirmation by NMR and biological testing. For biocatalytic synthesis of phase I metabolites, in the Expertise Bioreactions of Novartis Pharma AG, Basel (Switzerland), 14 different recombinant human (rh) cytochrome P450 isoenzymes functionally coexpressed with human P450 reductase in E. coli are established as whole cell biotransformation systems. The genetic constructs were established at the Biomedical Research Centre of the University of Dundee, Scotland, UK in the course of an academia/ industry collaboration between the said institute and 14 pharmaceutical companies. In a first step, often only the six major human CYP systems (CYP 1A2, 2C9, 2C19, 2D6, 3A4, 2E1) or otherwise, all 14 CYP-expressing E. coli cell lines available to us are screened to identify the relevant isoenzyme. This is then used as the biocatalyst for metabolite production via whole cell biotransformation. If the desired compound is not formed by human CYPs or the conversion is too low, a suitable microorganism can often be found producing the desired structure in high yield. For this purpose, a screening of 150 microbial strains with known oxidative capabilities is carried out in a 96-well format within 8 days (System Duetz, Kühner AG, Basel, Switzerland). The technology for preparative biotransformation involving CYPs is demonstrated below, showing the production of four metabolites from AAG561 (see Table 10.3), a new drug compound in development at Novartis Pharma AG against anxiety and depression. For completion, the latest developments of the methodology are given. 10.3.3.2 Production of E. coli Cells with CYP Activity From frozen glycerol stocks (−80 ºC, see below) of E. coli strains coexpressing CYPs and P450 reductase, some material was streaked onto a Luria Bertani (LB) agar plate containing 100 μg mL −1 ampicillin or 100 μg mL −1 of ampicillin plus 50 μg mL −1 of chloramphenicol, depending on the construction system used (both antibiotics for the systems with CYPs 1A2, 2C9, 2D6, and 3A5, where the two human enzymes are cloned into two plasmids, just ampicillin for the one plasmid systems with CYPs 1A1, 2C18, 2C19, 3A4, and 2E1). After incubation at 37 ºC for 16 h, a single colony was used to inoculate the preculture consisting of 25 mL solution of LB broth (Fluka, Switzerland) with pH 6.8 in a 100 mL shake flask contain-

10.3 Use of E. coli P450 Expression Systems in Bioreactors Table 10.3 Proposed structures and relative abundance of

metabolites from AAG561 after whole cell biotransformation with E. coli JM109 coexpressing CYP3A4 and P450 reductase.

NH

NH

N N N Cl

N Cl

NH N

HN

N

N

OH N Cl

NH N

N

NH

OH N Cl

N

NH N N

N

N

N

M4

AAG561

M3

M2

MH m/z 411,

MH m/z 357,

MH m/z 373,

MH+ m/z 385,

rel. peak area 62.9%

rel. peak area 9.2%

rel. peak area 2.1%

rel. peak area 2.9%

tR 15.4 min

tR 13.0 min

tR 11.24 min

tR 11.6 min

+

NH

+

+

N

N

N

N Cl

NH N

No structure proposal,

N

HN

N Cl

OH N Cl

NH N

NH N

molecule contains chloro atom

N

N

N

M7

M6

M5

M4 MH m/z 357,

MH m/z 369,

MH /z 427,

MH+ m/z 425,

rel. peak area 9.2%

rel. peak area 15.7%

rel. peak area 1.1%

rel. peak area 2.8%

tR 13.0 min

tR 13.24 min

tR 13.8 min

tR 15.2 min

+

+

+

ing antibiotics as mentioned above. The preculture was placed in an orbital shaker set at 37 ºC and a rate of 220 rpm until it reached an optical density at 600 nm (OD600 ) between 0.7 and 1. Then it was stored in a refrigerator until use (4 ºC, 1–2 days maximum). Main cultures were prepared with 100–200 mL of Modified Terrific Broth (MTB, Sigma-Aldrich, Buchs, Switzerland) in 500 mL flasks with 1–4 baffles (all combinations serve). The inoculation volume was 1% v/v. The flasks were incubated in an orbital shaker at 37 ºC and 180 rpm. Before induction of enzyme expression, the temperature was reduced to 30 ºC. Induction was performed at an OD600 of 0.7–1 by addition of 1 mmol L −1 of IPTG and 0.5 mmol L −1 of δaminolevulinic acid. The total cultivation time was around 24 h, in which all cell lines reached an OD600 of 14–16. The cells were centrifuged at 5000 rpm and 4 ºC for 15 min. The pellet was resuspended in one tenth of the original main

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culture volume using PSE buffer (KH2PO4/NaOH pH 7.2 50 mmol L −1, sucrose 250 mmol L −1, EDTA 0.25 mmol L −1) and, if not immediately applied, stored at −80 ºC until use as biocatalyst. For long-term preservation, precultures with an OD600 of 0.7–1 were frozen in the presence of 15% sterile glycerol at −80 ºC. On preparative scale, 14 × 500 mL Erlenmeyer flasks each fi lled with 200 mL of MTB medium plus 100 mg L −1 ampicillin were inoculated with E. coli JM109 coexpressing CYP3A4 and P450 reductase and further processed as described above. 10.3.3.3 Whole Cell Biotransformation In order to permeabilize the cells for biotransformation, 5% v/v of a Tween 80 solution (20 g L −1) was mixed into fresh or defrosted 10× cell suspension. After incubation for 4 min at room temperature, the cells were harvested by centrifugation (see above) resuspended in PSE buffer, centrifuged again and finally resuspended in this buffer (cells 10-fold concentrated compared to the original culture volume). Biotransformations on an analytical scale serve as activity assay for freshly grown cells and for optimizing the reaction conditions before preparative application. Furthermore, they are used for the identification of the CYP isoforms relevant for a specific compound and for the elaboration of its metabolite profi le. 0.5 mL of cell suspension were mixed with 2.5 μL of a methanolic solution of AAG561 (20 mg mL −1) in an Eppendorf cap closed with a membrane lid (Eppendorf Lid Bac, Eppendorf, Hamburg, Germany) and incubated at 30 ºC and 1100 rpm in a temperature-controlled Eppendorf mixer for 4 and 20 h. The reactions were stopped by mixing with 0.5 mL of acetonitrile for 15 min. After centrifugation in a Heraeus Biofuge at 14 000 rpm, the supernatant was subjected to RP18 -HPLC-DAD and RP18 -HPLC-MS analysis. The analytical biotransformations (4 h incubation time) showed metabolite formation with >10% conversion (rel. HPLC-MS peak area of the most abundant metabolite) by CYPs 1A1, 2C18, 3A4, and 3A5. The reactions involved were oxidative dealkylation and hydroxylation (see Table 10.3), whereby the dealkylated products M4 and M5 were the major reaction products. The additionally hydroxylated derivatives M2 and M3 as well as the di-dealkylated species M1 represented only minor metabolites. The data indicated that dealkylation preceded hydroxylation in the metabolic pathway. Since CYP3A4 showed the broadest metabolite pattern, the preparative biotransformation was performed with 280 mL of a 10× concentrated cell suspension of E. coli JM109 coexpressing this CYP isoform and P450 reductase after treatment with Tween 80 as described above. AAG561 was added in form of 1.4 mL of a methanolic solution (20 mg mL −1) and the incubation was performed in two 500 mL Erlenmeyer flasks with one baffle at 30 ºC and 180 rpm in a laboratory shaker for 20 h. The biotransformation broth was extracted twice with 200 mL of CH2Cl2/isopropanol (9 : 1). Phase separation was achieved by centrifugation. The organic layers were pooled, dried with MgSO4 and the solvent was removed under reduced pressure yielding ~360 mg of solid. Further purification was effected by prep. RP18 -HPLC (water–acetonitrile gradient in the presence of 0.05% trifluoro-


acetic acid) with UV and MS detection. All preparations obtained thereby still contained some Tween 80 originating from the cell treatment for permeabilization. These contaminations, however, could be removed by normal phase chromatography. The structures of the preparations of M1–M5 were elucidated by NMR spectroscopy and the identity and purity was verified by HPLC-MS (see above) and, except for M3, by HPLC-UV (220 nm). Up to 13.6 mg of the different metabolites were obtained with purities between 74 and 98%. Metabolites M2 and M3 proved to be hydroxylated at the methyl group on the quinoxaline ring (see Fig. 10.3). The metabolite pattern in respect to the structure and even the quantity of the reaction products was identical to that from in vitro incubations of [14C]AAG561 with human postmitochondrial liver fractions (S9) measured by HPLC with radiodetection (Wirz B and Weber B, internal report, Novartis Basel, Switzerland).

Fig. 10.3 Metabolites prepared from AAG561 via whole cell biotransformation using CYP3A4/P450 reductase coexpressed in E. coli JM109.

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10.3.3.4 Recent Developments E. coli cells with high activity of CYPs can also be obtained on larger scale by cultivation in a stainless steel stirred tank bioreactor. The aeration should be sufficient for good growth but during CYP expression the dissolved oxygen concentration should be kept very low in order to avoid enzyme deactivation [42]. Recently we have grown the cells on a routine basis in a BioWave 50SPS bioreactor (WaveBiotech AG, Tagelswangen, Switzerland) in order to lower the costs by use of a cheaper and simpler technology. A disposable polyethylene bag serves as the reactor, which is rocked on a temperature-controlled table. Oxygen is supplied via a stream of sterile air through the headspace of the bag. Under conditions recommended for the cultivation of E. coli (40 rocks/min, 10.5º rocking angle) and an airflow of 0.5 L min−1 (maximum value of the built-in membrane pump), the CYP3A4-expressing E. coli cell line grew only to an OD600 of 8 instead of 16 commonly achieved in MTB, probably because of oxygen limitation. By supplying an air flow supplemented with 10% of pure oxygen, cell density and CYP activity are identical to the ones obtained in shake flasks. The treatment of the biocatalytically active cells with the Tween 80 implicated an additional purification step for separating off the detergent. Alternatively, substrates with low water solubility can be coated onto highly dispersed silica gel before supplying to the reaction mixture. The considerable increase in conversion is probably due to the enhanced dissolution via offering a large solid–liquid interface. Applying these more recent technologies, a compound of undisclosed structure with a molecular weight (MW) of 533 and a calculated logP of 2.29 (program CPOGP, BioByte Corp., Claremont, CA, USA) could be converted to 80% under catalysis of CYP3A4 to 100 mg of a N-demethylated metabolite, another compound (MW 241, calc. logP 2.56) by CYP2D6 even in a concentration of 200 mg L −1 to 100% to a monohydroxylated derivative (300 mg). In comparison it can be assumed that the relatively low conversion of AAG561 (see Table 10.3) was probably, at least to some extent, due to its high hydrophobicity (calc. logP 6.9) and hence, low water solubility.

10.4 Conclusion

E. coli systems expressing human P450s have been used on a small scale in preclinical drug development (e.g. metabolic high-throughput screening). However, we have also convincingly demonstrated the effectiveness of E. coli expressing human CYP450 isozymes as biocatalysts for the production of drug metabolites on the 100 mg to 1 g scale. For some purposes it may be advantageous to employ broken cells, while for others intact cells may be preferable. Often such metabolites are either not accessible via chemical synthesis or only via very tedious multistep procedures. The biocatalytic route offers a convenient, relatively fast preparation of drug metabolites for a variety of applications. The availability

References

of 14 cloned human enzymes means that the authentic metabolites found in vivo will be accessible in the vast majority of cases. In order to fully exploit the potential of these enzymes the respective fermentation and downstream processing capacities should be available.

Acknowledgments

The following investigators at Roche are acknowledged: We would especially like to mention Dr Peter Matzinger who established the methods for fermentation of the LINK strains and who demonstrated the value of these reagents as biocatalysts. We also greatly acknowledge Dr Ernst Kupfer for downstream processing and metabolite purification and Marie-Odile Grieneisen, Jacky Joerger, and Marie-Joelle Sikora for technical support. I also thank Dr Wolfgang Weber for allowing use of the fermentation facility and Dr Beat Wirz and Dr Hans Iding for helpful discussions. The following staff at Novartis are acknowledged: We are grateful to Dr J. Nozulak for providing AAG561 and M4 as reference compound, Dr L. Oberer for NMR spectroscopy and to Mr A. Meishammer and Dr E. Francotte for purification of the metabolites. The following researchers at the Biomedical Research Centre (BMRC) in Dundee are acknowledged: We thank Drs Mike Voice and Mike Pritchard (formerly BMRC, now CYPEX limited), Lesley McLaughlin, and Dongtao Li for their skilful work. The program at the BMRC was sponsored by the Biological Sciences Research Council, the UK Department of Trade and Industry and the LINK consortium of pharmaceutical companies: Astra, Glaxo-Wellcome, Janssen Pharmaceutica, Lilly, Novo-Nordisk, Park-Davis, Pfizer, Hoffman LaRoche, Sanofi-Winthrop, Servier, Smith-Kline Beecham, Wyeth-Ayerst and Zeneca. Their generous support and their helpful discussions are gratefully acknowledged.

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16 Kemp, C.A., Flanagan, J.U., van Eldik, A.J., et al. Validation of model of cytochrome p450 2D6, An in silico tool for predicting metabolism and inhibition. J Med Chem 2004, 47, 5340–5346. 17 Friedberg, T., Pritchard, M.P., Bandera, M., et al. Merits and limitations of recombinant models for the study of human P450-mediated drug metabolism and toxicity: An intralaboratory comparison. Drug Metab Rev 1999, 31, 523–544. 18 Guengerich, F.P. Cytochrome P450 enzymes in the generation of commercial products. Nat Rev Drug Discov 2002, 1, 359–566. 19 Smith, D.A., Obach, R.S. Seeing through the mist: Abundance versus percentage. Commentary on Metabolites in safety testing. Drug Metab Dispos 2005, 33, 1409–1110. 20 Gold, L. Expression of heterologous proteins in Escherichia coli. In: Goeddel, D.V., ed. Methods in Enzymology. San Diego: Academic Press, 1990, pp. 11– 310. 21 Blake, J.A.R., Pritchard, M., Ding, S., et al. Coexpression of a human P450 (CYP3A4) and P450 reductase generates a highly functional monooxygenase system in Escherichia coli. FEBS Lett 1996, 397, 210–214. 22 Pritchard, M.P., Glancey, M.J., Blake, J.A.R., et al. Functional coexpression of CYP2D6 and human cytochrome P450 reductase in Escherichia coli. Pharmacogenetics 1998, 8, 33–42. 23 Pritchard, M.P., Ossetian, R., Li, D.N., et al. A general strategy for the expression of recombinant human cytochrome P450s in Escherichia coli using bacterial signal peptides: Expression of CYP3A4, CYP2A6, and CYP2E1. Arch Biochem Biophys 1997, 345, 342–354. 24 Pritchard, M.P., McLaughlin, L.A., Friedberg, T. Establishment of functional human cytochrome P450 monooxygenase systems in Escherichia coli. In: Phillips, I. R., Shephard, E.A., eds. Cytochrome P450 Protocols. Totowa, NJ: Humana Press, 2006, pp. 19–29.

References 25 Campbell, I., Duffus, J.H., eds. Yeast: A Practical Approach. Washington: IRL Press, 1988. 26 Emr, S.D. Heterologous gene expression in yeast. In: Goeddel, D.V., ed. Methods in Enzymology. San Diego: Academic Press, 1990, pp. 231–279. 27 Urban, P., Truan, G., Bellamine, A., Laine, R., Gautier, J.C., Pompon, D. Engineered yeast simulating P450dependent metabolism: Tricks, myths and reality. In: Kingsley N, ed. Drug Metabolism and Interactions. London: Freund Publishing, 1994, pp. 169–200. 28 Shkumatov, V.M., Usova, E.V., Radyuk, V.G., et al. Oxidation of 17 alpha,20 beta- and 17 alpha,20 alphadihydroxypregn-4-en-3-ones, side products of progesterone biotransformation with recombinant microorganisms expressing cytochrome P-45017 alpha. Russian J Bioorg Chem 2003, 29, 581–610. 29 Nthangeni, M.B., Urban, P., Pompon, D., Smit, M.S., Nicaud, J.M. The use of Yarrowia lipolytica for the expression of human cytochrome P450CYP1A1. Yeast 2004, 21, 583–592. 30 Dietrich, M., Grundmann, L., Kurr, K., et al. Recombinant production of human microsomal cytochrome P450 2D6 in the methylotrophic yeast Pichia pastoris. Chembiochem 2005, 6, 2014–2022. 31 Cullin, C., Pompon, D. Synthesis of functional cytochromes P450 P1 and chimeric P-450 P3–1 in the yeast Saccharomyces cerevisiae. Gene 1988, 65, 203–210. 32 Bligh, H.F.J., Wolf, C.R., Smith, G., Beggs, J.D. Production of cytochrome P450 reductase yeast-rat hybrid proteins in Saccharomyces cerevisiae. Gene 1992, 110, 33–39. 33 Pompon, D., Louerat, B., Bronine, A., Urban, P. Yeast expression of animal and plant P450s in optimized redox environments. In: Johnson, E.F., Waterman, M.R., eds. Methods in Enzymology. San Diego: Academic Press, 1996. pp. 51–64. 34 Paine, M.J.I., Gilham, D., Roberts, G.C.K., Wolf, C.R. Functional high-level

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11 Human Cytochrome P450 Monooxygenases – a General Model of Substrate Specificity and Regioselectivity Jürgen Pleiss 11.1 Introduction

Cytochrome P450 monooxygenases (CYPs) constitute a large, ubiquitous enzyme family. They catalyze a wide range of reactions such as hydroxylation of aromatic or aliphatic compounds, epoxidation, N-demethylation, O-demethylation, and exhibit a broad substrate specificity. They are the most important drug-metabolizing enzymes, being responsible for 75% of phase I-dependent drug metabolism [1]. The human genome encodes 57 putatively functional CYP genes and 58 pseudogenes [2]. The major isoforms of importance for drug metabolism are CYP2C9, CYP2C19, CYP2D6, and CYP3A4 [1]. In addition, a rapidly increasing number of polymorphisms have been identified that lead to inactivation of the respective enzyme variant. In many cases, these genetic factors have a major impact on the pharmacokinetics and pharmacodynamics of a particular drug and thereby influence the sensitivity in an individual patient [3]. Currently the sequences of more than 4000 CYPs are known. A systematic nomenclature scheme exists which mainly depends on sequence similarity [4]. CYPs have been assigned to a hierarchy of protein families based on sequence similarity: Proteins within a given superfamily have a sequence identity >40%, proteins within a homologous family >55%. Between superfamilies, overall sequence similarity is low, with the exception of a few highly conserved sequence patterns such as the ExxR motif in helix K [5] or the heme-binding motif GxRxCxG. Despite their differences in sequence and substrate specificity, the structures of CYPs are highly similar, as can be concluded from 106 X-ray structures of 25 different CYPs. In all CYPs the catalytically active heme is buried deeply inside the protein. The substrate-binding site is a large cavity that is connected to the heme by a narrow channel but shielded from the solvent in most X-ray structures. For most CYPs several X-ray structures are available which differ due to different crystallization conditions or due to binding of a substrate. In many cases local conformational changes led to an opening of channels from the solvent to the substrate-binding site.

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11 Human Cytochrome P450 Monooxygenases

Fig. 11.1 Geometry (distances and angles in Ångstroms and degrees, respectively) of the reactive complex of compound I and substrate [9].

The reaction mechanism has been investigated by spectroscopic [6, 7] and quantum chemical methods [8]. In the reactive state the substrate has to approach the heme iron at a well-defined geometry [9] with the distance between the hydrogen of the aliphatic C-H group and the heme oxygen of 2.2 Å and the C-H-O angle of 175 degrees (Fig. 11.1). Since the reactive site of the substrate has to be near to the catalytic iron, the shape of the protein and its flexibility are expected to determine its activity and regioselectivity towards a given substrate. Thus, computational approaches for structure-based drug design have been successfully applied to identify putative substrates of human CYPs from large libraries of molecules [10, 11] or to investigate the mechanism of inhibition [12]. However, all docking approaches assume a rigid substrate-binding site and therefore the success rate of GRID-based methods or molecular docking approaches is still limited for enzymes with flexible substrate-binding sites. To learn more about the molecular basis of the biochemical properties of CYPs it is promising to combine different methods: the systematic comparison of sequence and structure of all CYPs and the molecular modeling of the dynamics of protein–substrate complexes. The questions to address are: • How can the effect of non-synonymous single nucleotide polymorphisms to activity be predicted? • Can regioselectivity be predicted by comparing the shape of the binding site? • How does the substrate get into the binding site? • What is the role of flexibility in regioselectivity? 11.2 What Can We Learn From Sequence? 11.2.1 The Cytochrome P450 Engineering Database (CYPED)

The data warehouse system DWARF (Data Warehouse ARchitecture for protein classiFication) has been set up to systematically analyze sequences and structures

11.2 What Can We Learn From Sequence?

Fig. 11.2 Modular concept of the DWARF system.

of protein families and to study the relationship between sequence, structure, and function of enzymes [13]. The DWARF system consists of a relational database, analysis tools, a so-called ETL layer (Extraction, Transformation, Loading) to retrieve data from public resources, and a set of tools to analyze sequence and structure data (Fig. 11.2). The ETL layer parses sequence data from public databases; the analysis tools are used to assign proteins to homologous families and superfamilies based on multisequence alignments, to annotate amino acid positions, and to manually enrich information. The database is then published on a web server. Recently this system has been applied to establish the Cytochrome P450 Engineering Database (CYPED). The creation of this database was based on seed sequences from the Cytochrome P450 Homepage [14]. Sequence data is extracted from GenBank [15] and SWISS-PROT [16], structure data from the Protein Data Bank (PDB [17]). In a further clustering stage the proteins were assigned based on sequence similarity and according to Nelson’s classification scheme [4] to 550 superfamilies and 1100 homologous families. The functions implemented in DWARF were applied to functionally annotate sequences and automatically transfer these annotations on other family members. The annotation was validated and enriched by multisequence alignments, superimposed structures and phylogenetic analysis (Fig. 11.3). The final database is published online and can be browsed either by families, organisms, structure, or systematic CYP nomenclature. Alternatively, a BLAST search interface makes it possible to search for similar sequences. Automatic update tools assure that CYPED keeps pace with progress in the field of CYPs. Apart from the CYPED, two family databases have been published previously: the Lipase Engineering Database [18] and a smaller database on glycoside hydrolases [19]. 11.2.2 The Effect of Mutations on Activity

The interindividual and interethnic variability in xenobiotic metabolism and drug responses is a significant clinical challenge. Because the majority of phase Idependent drug metabolism is carried out by polymorphic enzymes, a drug

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Fig. 11.3 Creation, clustering, publishing and update of the database.

level in plasma can vary more than 1000-fold between two individuals having the same weight and with the same drug dosage [20]. The cytochrome P450 enzymes in family 1–3 are generally polymorphic. Information on alleles and their effect on enzyme expression or function can be retrieved from the homepage of the Human Cytochrome P450 Allele Nomenclature Committee (http://www.cypalleles.ki.se/). However, for many of the alleles no information is available about the effect on enzyme activity. Clearly it would be helpful to predict whether a non-synonymous variation is tolerated or whether it leads to a loss of function [21]. It has been shown that diseaseassociated mutations are conserved evolutionarily [22, 23] and that they are relevant to protein structure [24]. The first studies to predict the effect of mutations quantified the extent of changes by a amino acid scoring function such as BLOSUM62 [25]. Later methods took the sequence context of the mutation into account. The SIFT method [16, 22] is based on a position-specific scoring matrix and analyzes the variability of the mutated position. Other methods include protein structural features such as solvent accessibility, secondary structure, and disulfide bridge formation [24]. Decision trees and support vector machines (SVMs) have also been applied to analysis based on structural and evolutionary properties [26]. A systematic sequence analysis has been used to predict the effect of single point mutations in human cytochrome P450 monooxygenases. Known single nucleotide polymorphisms were retrieved from the homepage of the Human CYP Allele Nomenclature Committee [27]. For non-synonymous polymorphisms that led to an amino acid substitution, their effects on enzymatic activity were correlated with the conservation of the respective position in a multiple sequence

11.2 What Can We Learn From Sequence?

alignment of the homologous family. Conservation scores were calculated using the program al2co [28] and the unweighted sum of pairs method with the BLOSUM62 scoring matrix. A simple rule was derived: mutations at highly conserved positions led to inactivation, mutations at variable positions were neutral. In the CYP2D family, position 324 is highly conserved with a conservation score of 6 (Table 11.1). Exchange of histidine by proline leads to inactivation of the enzyme. In contrast, position 437 is variable with a conservation score of 1.2. Here an exchange of alanine to serine is neutral. For six alleles of 2D6 the effect of the polymorphism could be predicted by conservation score (Fig. 11.4): mutations in Table 11.1 Analysis of CYP2D6 alleles with known activity data from the human CYP allele homepage. For alleles with more than one amino acid exchange the conservation score from the amino acid position with the highest conservation score was used.

Allele

Amino acid exchange

Conservation score

Activity

CYP2D6*2 CYP2D6*7 CYP2D6*10A CYP2D6*14A

R296C; S486T H324P P34S; S486T P34S; G169R; R296C; S486T A437S V11M; R296C; S486T

2.5 6.0 7.0 7.0

Increased No activity Decreased No activity

1.2 3.0

Neutral Neutral

CYP2D6*33 CYP2D6*35

Fig. 11.4 Correlation of experimental enzyme activity and amino acid conservation calculated from the multisequence alignment of the CYP2D family (Table 11.1).

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the variable positions 11, 296, and 437 are neutral or even led to an increased activity, mutations in the conserved positions 34 and 324 led to a reduced activity or even loss of activity. A similar approach has been applied to analyze the effect of polymorphisms in the murine 21-hydroxylase gene (Cyp21). The combination of sequence conservation analysis and three-dimensional structure modeling provides a valuable tool to understand the effects of the individual mutations and polymorphisms on the resulting enzyme activity. The general effect of the mutations on activity can be explained by a conservation score. Inactivating mutations were either found near the reductase-binding site or in the substrate-binding pocket. Thus three classes of mutations could be assigned: neutral mutations, mutations with a loss of enzyme activity caused by inefficient electron flux, and mutations with a loss of activity due to deficient substrate binding [29].

11.3 What Can We Learn from Structure? 11.3.1 The Role of Flexibility

To model the effect of structure and flexibility to activity and selectivity, we investigated P450-catalyzed hydroxylation of the drug warfarin. Warfarin has multiple sites that can be hydroxylated. An estimation of chemical reactivity by the PETRA method [30] predicted position 10 to be most reactive, positions 4, 6, and 7 being slightly less reactive. Thus, hydroxylation by enzymes that lack regioselectivity should result in a mixture of 10-, 4-, 6-, and 7-hydroxywarfarin. CYP2C9, however, is highly selective and prefers positions 7 and 6 [31]. In contrast, CYP3A4 is complementary to CYP2C9 and hydroxylates preferentially in positions 4 and 10 [32]. Thus, regioselectivity is encoded in the structure and the dynamics of these enzymes. Recently, an X-ray structure of CYP2C9 in complex with (S)-warfarin has been published [33]. The substrate-binding site is a large cavity inside the protein with a narrow channel leading to the catalytically active heme, but no channel that is large enough for the substrate leading to the outside. The substrate binds in a nonproductive position far away from the heme. Thus, flexibility of the protein is prerequisite to its activity: for the substrate to enter the binding cavity, then to approach the heme through a narrow heme access channel, and finally for the product to leave the binding site. A series of molecular dynamics simulations of the CYP2C9– warfarin complex in water demonstrated the high degree of flexibility of this protein [34]. The major motions were found in the B and C and the F and G helices. Helices F and G are part of the binding cavity, the loops between helices B and C and between helices F and G are blocking the binding cavity from the solvent. In the X-ray structure narrow channels between helix F and the C-terminal antiparallel β-sheet, and between helix B and the loop between helix F and G lead

11.3 What Can We Learn from Structure?

from the solvent to the binding site, but they are too narrow for bulky substrate or product molecules to enter or leave the binding site. However, in the course of molecular dynamics simulations the warfarin molecule bound into each of the channels and stabilized them in a widely open conformation. The occurrence of temporary channels has been observed in many CYPs. In a systematic analysis of all currently available X-ray structures a multitude of possible substrate channels in mammalian and bacterial CYPs have been identified [35]. It has been proposed that there might be different channels for hydrophobic and hydrophilic substrates [36]. Sixteen different simulations starting with the experimentally determined, non-productive CYP2C9–warfarin complex were performed [34]. In 12 simulations, the substrate stayed near its position in the X-ray structure, while in three simulations it moved into a different, non-productive state. However, in one simulation the substrate moved toward the heme from its non-productive position along the narrow heme access channel. In its final orientation, it pointed toward the heme with its 6 and 7 positions. Most frequently the hydrogen at position 7 was closer than 3 Å to the oxyferryl group, less frequently the hydrogen at position 6. This preferred substrate orientation in the narrow heme access channel is consistent with the experimentally observed regioselectivity, resulting in 71% 7hydroxywarfarin and 22% 6-hydroxywarfarin. 11.3.2 The Role of Binding Site Shape

Dextromethorphan (Fig. 11.5) is a widely used antitussive drug which is metabolized in the liver. Two CYPs are mainly involved in demethylation: CYP2D6 which preferentially catalyzes O-demethylation and CYP3A4 which catalyzes N-demethylation in vitro [37]. Three factors might contribute to this regioselectivity: the chemical reactivity of the two sites, the shape of the substrate, and the shape of the binding site.

Fig. 11.5 Structure of dextrometorphan.

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The chemical reactivities of the two sites in dextromethorphan were predicted by the PETRA method [30]. By comparing the bond dissociation energy, PETRA predicted a higher reactivity of N- as compared to O-demethylation. The two sites also differ in accessibility: the O-position is more accessible than the N-position. Thus, the substrate is characterized by two reactive ends: the Omethyl group that is less reactive but accessible, and the N-methyl group that is more reactive and located at the bulky end of the molecule. As a consequence, N-demethylation is preferred by enzymes that provide unrestricted access to the active site, while only O-demethylation can occur if the access to the active site is narrow. By comparing the shape of the substrate-binding sites, CYP2D6 and CYP3A4 can be assigned to either of two classes, depending on the width of their heme access channel (Fig. 11.6). CYP2D6 belongs to the class of CYPs with a narrow access channel, and the substrate is expected to preferentially bind to the heme with its accessible site. As a consequence, this model predicts that CYP2D6 preferentially catalyzes O-methylation. In contrast, CYP3A4 is expected to accept both sites due to its wide access channel. As a consequence, the more reactive group is expected to be demethylated preferentially, thus N-demethylation is predicted as observed experimentally. A similar analysis has recently been extended to two additional drugs, verapamil and propafenone. Both molecules have two competing sites which are selectively modified by CYP2C9 and CYP3A4. As in dextromethorphan, the experimentally observed regioselectivity was predicted by a simple rule: the more accessible but less reactive site is preferred by CYP2C9, which has a narrow access channel to heme, while the more reactive but less accessible site is preferred by CYP3A4.

Fig. 11.6 Preferred orientation of dextromethorphan in the binding sites of CYP2D6 (left) and CYP3A4 (right). Therefore, the less reactive but more accessible O-methyl group (blue) and the chemically reactive N-methyl group (red) are preferred by CYP2D6 and CYP3A4, respectively.

11.4 Conclusion

11.4 Conclusion

The family of CYPs is characterized by the conserved structure of a rigid protein core, high structural mobility, and sequence variability of the residues forming the substrate-binding site, low global sequence similarities between different superfamilies, and a high diversity of substrates and products. To derive a general model of substrate specificity and regioselectivity for human CYPs, a systematic comparison of sequence and structure, and a simulation of protein dynamics were performed. A set of simple rules could be derived: 1. Enzymatic activity conferred by non-synonymous single nucleotide polymorphisms can be predicted by analyzing the conservation on amino acid level inside a homologous family (e.g. CYP2D). Exchange of a residue with high conservation score will probably lead to a variant with decreased activity. High conservation is indicative of residues that are essential to substrate binding, electron transport, or overall protein stability. Interestingly, the role of individual residues might be family-specific. Therefore, the conservation analysis should be limited to homologous families. 2. The substrate-binding site consists of a large, mobile pocket, and a narrow, rigid access channel that leads to the catalytic heme. Temporary channels which are family-, substrate-, and product-specific connect the binding pocket to the solvent. This delicate balance between rigidity and mobility of the CYP–substrate complex accounts for the observed biochemical properties that are experimentally observed for most mammalian CYPs: broad substrate specificity and high regioselectivity. Due to the size and flexibility of the substrate-binding site, the substrate can bind in many states. Only a small number of states are expected to be productive, while most of these states are non-productive. 3. For substrates with two reactive sites, three factors contribute to regioselectivity: the shape of the binding site, the shape of the substrate, and the relative chemical reactivity of the two reactive sites. CYPs can be classified by the width of their heme access channel. Enzymes with a narrow channel such as CYP2D6 prefer the site that has the highest accessibility, while enzymes with a wide access channel such as CYP3A4 prefer the site with highest chemical reactivity.

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Acknowledgments

Valuable contributions by Markus Fischer, Michael Knoll, Michael Krahn, Alexander Seifert, Demet Sirim, and Stephan Tatzel are gratefully acknowledged. This work was supported by the German Research Foundation and the German Federal Ministry of Education and Research.

References 1 Ingelman-Sundberg, M. Human drug metabolising cytochrome P450 enzymes: properties and polymorphisms. Naunyn Schmiedebergs Arch Pharmacol 2004, 369, 89. 2 Nelson, D.R., Zeldin, D.C., Hoffman, S.M., Maltais, L.J., Wain, H.M., Nebert, D.W. Comparison of cytochrome P450 (CYP) genes from the mouse and human genomes, including nomenclature recommendations for genes, pseudogenes and alternative-splice variants. Pharmacogenetics 2004, 14, 1. 3 Oscarson, M. Pharmacogenetics of drug metabolising enzymes: importance for personalised medicine. Clin Chem Lab Med 2003, 41, 573. 4 Nelson, D.R., Koymans, L., Kamataki, T., Stegeman, J.J., Feyereisen, R., Waxman, D.J., Waterman, M.R., Gotoh, O., Coon, M.J., Estabrook, R.W., Gunsalus, I.C., Nebert, D.W. P450 superfamily: update on new sequences, gene mapping, accession numbers and nomenclature. Pharmacogenetics 1996, 6, 1. 5 Hasemann, C.A., Kurumbail, R.G., Boddupalli, S.S., Peterson, J.A., Deisenhofer, J. Structure and function of cytochromes P450: a comparative analysis of three crystal structures. Structure 1995, 3, 41. 6 Davydov, R., Makris, T.M., Kofman, V., Werst, D.E., Sligar, S.G., Hoffman, B.M. Hydroxylation of camphor by reduced oxy-cytochrome P450cam: mechanistic implications of EPR and ENDOR studies of catalytic intermediates in native and mutant enzymes. J Am Chem Soc 2001, 123, 1403. 7 Thomann, H., Bernardo, M., Goldfarb, D., Kroneck, P.M.H., Ullrich, V.

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Evidence for water binding to the Fe center in cytochrome P450Cam obtained by 17O Electron Spin Echo Envelope modulation spectroscopy. J Am Chem Soc 1995, 117, 8243. Shaik, S., Kumar, D., de Visser, S.P., Altun, A., Thiel, W. Theoretical perspective on the structure and mechanism of cytochrome P450 enzymes. Am Chem Soc 2005, 105, 2279. Schoneboom, J.C., Cohen, S., Lin, H., Shaik, S., Thiel, W. Quantum mechanical/molecular mechanical investigation of the mechanism of C-H hydroxylation of camphor by cytochrome P450cam: theory supports a two-state rebound mechanism. J Am Chem Soc 2004, 126, 4017. Cruciani, G., Carosati, E., De Boeck, B., Ethirajulu, K., Mackie, C., Howe, T., Vianello, R. MetaSite: understanding metabolism in human cytochromes from the perspective of the chemist. J Med Chem 2005, 48, 6970. de Graaf, C., Oostenbrink, C., Keizers, P.H., van der Wijst, T., Jongejan, A., Vermeulen, N.P. Catalytic site prediction and virtual screening of cytochrome P450 2D6 substrates by consideration of water and rescoring in automated docking. J Med Chem 2006, 49, 2417. Richter, T., Murdter, T.E., Heinkele, G., Pleiss, J., Tatzel, S., Schwab, M., Eichelbaum, M., Zanger, U.M. Potent mechanism-based inhibition of human CYP2B6 by clopidogrel and ticlopidine. J Pharmacol Exp Ther 2004, 308, 189. Fischer, M., Thai, Q.K., Grib, M., Pleiss, J. DWARF – a data warehouse system for analyzing protein families. BMC Bioinformatics 2006, 7, 495.

References 14 Nelson, D.R. Mining databases for cytochrome P450 genes. Methods Enzymol 2002, 357, 3. 15 Benson, D.A., Karsch-Mizrachi, I., Lipman, D.J., Ostell, J., Wheeler, D.L. GenBank. Nucleic Acids Res 2006, 34, D16. 16 Boeckmann, B., Bairoch, A., Apweiler, R., Blatter, M.C., Estreicher, A., Gasteiger, E., Martin, M.J., Michoud, K., O’Donovan, C., Phan, I., Pilbout, S., Schneider, M. The SWISS-PROT protein knowledgebase and its supplement TrEMBL in 2003. Nucleic Acids Res 2003, 31, 365. 17 Berman, H.M., Westbrook, J., Feng, Z., Gilliland, G., Bhat, T.N., Weissig, H., Shindyalov, I.N., Bourne, P.E. The Protein Data Bank. Nucleic Acids Res 2000, 28, 235. 18 Fischer, M., Pleiss, J. The Lipase Engineering Database: a navigation and analysis tool for protein families. Nucleic Acids Res 2003, 31, 319. 19 Strohmeier, M., Hrmova, M., Fischer, M., Harvey, A.J., Fincher, G.B., Pleiss, J. Molecular modeling of family GH16 glycoside hydrolases: potential roles for xyloglucan transglucosylases/hydrolases in cell wall modification in the poaceae. Protein Sci 2004, 13, 3200. 20 Ingelman-Sundberg, M. Pharmacogenetics: an opportunity for a safer and more efficient pharmacotherapy. J Intern Med 2001, 250, 186. 21 Bowie, J.U., Reidhaar-Olson, J.F., Lim, W.A., Sauer, R.T. Deciphering the message in protein sequences: tolerance to amino acid substitutions. Science 1990, 247, 1306. 22 Chasman, D., Adams, R.M. Predicting the functional consequences of nonsynonymous single nucleotide polymorphisms: structure-based assessment of amino acid variation. J Mol Biol 2001, 307, 683. 23 Mooney, S.D., Klein, T.E. The functional importance of disease-associated mutation. BMC Bioinformatics 2002, 3, 24. 24 Sunyaev, S., Ramensky, V., Bork, P. Towards a structural basis of human non-

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36 Schleinkofer, K., Sudarko, Winn, P.J., Ludemann, S.K., Wade, R.C. Do mammalian cytochrome P450s show multiple ligand access pathways and ligand channelling? EMBO Rep 2005, 6, 584. 37 Yu, A., Haining, R.L. Comparative contribution to dextromethorphan metabolism by cytochrome P450 isoforms in vitro: can dextromethorphan be used as a dual probe for both CTP2D6 and CYP3A activities? Drug Metab Dispos 2001, 29, 1514–1520.

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12 Approaches to Recycling and Substituting NAD(P)H as a CYP Cofactor Dirk Holtmann and Jens Schrader 12.1 Introduction

A key issue in the use of isolated cytochrome P450 monooxygenases (P450s) in catalytic applications is the demand for a continuous electron supply by its prosthetic heme iron group. In vivo, the electrons are supplied by cellular NAD(P)H which is, however, far too expensive to be used in equimolar concentrations during technical applications [1]. Therefore, several approaches to substituting or recycling NAD(P)H continuously during biocatalytic reactions have been explored. A useful cofactor regeneration/substitution method has to fulfi l several requirements [2]. The enzymes, reagents, immobilization matrices, electrodes and further equipment required should be readily available, inexpensive, easy to manipulate, and stable under the operational conditions. No reagents or byproducts of the regeneration/substitution step should interfere with product isolation and they should be compatible with the rest of the reaction system. During their catalytic cycle P450s act with specific redox partners: mitochondrial systems include two proteins, NADPH-specific adrenodoxin reductase with FAD cofactor, and adrenodoxin, a 2Fe-2S ferredoxin-type iron–sulfur protein [3]. Microsomal systems depend on a single NADPH-specific P450 reductase that contains both FAD and FMN as cofactors. FAD can accept two electrons from NADPH and FMN functions as a single electron carrier. Some microsomal P450s may receive the second electron from NADH by cytochrome b5 reductase and cytochrome b5. CYP102A1 from Bacillus megaterium (P450BM-3) is catalytically self-sufficient and contains both the reductase and P450 domains on a single polypeptide chain. The two domains of P450BM-3 appear to be analogous in their function and homologous in their sequence to the microsomal P450 system components. Bacterial P450cam-type systems include a ferredoxin reductase and a ferredoxin (named putidaredoxin) that are functionally similar to the mitochondrial electron transfer proteins. CYP101 from Pseudomonas putida (P450cam) and the electron transfer proteins are all soluble and do not appear to be membrane associated. All these redox partners are possible targets for the regeneration or

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12 Approaches to Recycling and Substituting NAD(P)H as a CYP Cofactor

substitution of the natural electron transfer system. For a general overview of developments in the area of cofactor regeneration and substitution independent of P450 monooxygenases see [2, 4–7]. This review focuses on the application of the recycling or substitution systems of P450s for biooxidation. The applications of biosensing are only marginal aspects of this chapter, for instance when the general principles of the systems are necessary for understanding. For further reading about the application of the regenerating and substituting systems in the area of biosensing see [8–10].

12.2 Chemical Substitution of Cofactors

The direct regeneration of cytochrome P450 systems with dithionite has been reported [11, 12]. The ability of dithionite, an inexpensive reducing agent routinely used to produce the ferrous-carbonyl form of P450, to support P450BM-3catalyzed hydroxylation of palmitic acid was studied [11]. The hydroxylation products in the presence of dithionite were 15-, 14-, and 13-hydroxyhexadecanoate, with relative distributions similar to those observed with NADPH. The hydroxylation reaction was carried out in two separate steps: anaerobic reduction and subsequent oxidation of P450BM-3 by oxygen bubbling. The reduction step was much slower than the oxidation step, thus limiting the overall rate of hydroxylation. Upon addition of dithionite, the reductase domain of P450BM-3 seemed to be reduced before significant reduction of the heme domain occurred. In comparison with the rate of the P450BM-3 heme iron reduction by NADPH, the rate of reduction by dithionite is slow. It was shown that the direct chemical reduction of heme iron is very inefficient. During hydroxylation of palmitic acid by P450BM-3, for example, the reduction of oxygen in the presence of sodium dithionite was at least 8000 times lower than that by NADPH. A further approach to overcoming the problem of cofactor costs is the use of the peroxide shunt pathway with strong oxidants, such as hydrogen peroxide or periodate, which directly catalyze the reduction of the heme iron [13, 14]. The strong oxidants transfer one oxygen atom directly to the ferric/ferrous atom at the catalytic centre of the heme. The overall reaction equation using hydrogen peroxide is: RH + H2O2 → ROH + H2O This method is associated with the problem of fast inactivation of the enzyme due to heme degradation at the required peroxide concentrations, as indicated by a decrease in the heme absorbance peak in the presence of the oxidant. The hydrogen peroxide-driven reaction was investigated with some unmodified P450s (e.g the epoxidation of styrene) [15, 16] and reactions of CYP2B4 with aniline, aminopyrine and p-nitroanizole [17]. The two human P450 isoforms, CYP2D6

12.3 Enzymatic Regeneration of Cofactors

and CYP3A4, were selected by Chefson et al. because of their high substrate promiscuity, a significant advantage for use in synthesis [18]. Neither of these enzymes showed significant activity in the presence of aqueous hydrogen peroxide. It was shown that without mutagenesis, the two studied isoforms can use various hydrogen peroxide donors or organic peroxides, such as sodium percarbonate (SPC), cumene hydroperoxide (CHP), and tert-butylhydroperoxide (tBHP), to catalyze reactions in aqueous media. The best cofactor replacement was CHP, which, after 1 h, yielded up to 210% of the amount of product obtained under natural conditions. The mechanism of hydroxylation has been examined by employing NaIO4, NaClO4, and organic hydroperoxides as hydroxylating agents in steroid hydroxylation reactions catalyzed by partially purified cytochrome P450 as the catalyst [19].

12.3 Enzymatic Regeneration of Cofactors

A popular idea with regard to NAD(P)H regeneration is the use of a second enzyme such as a dehydrogenase in a coupled reaction [20, 21]. This approach has already led to a wider use of redox enzymes for chemical synthesis. The regeneration of the cofactor requires a second enzyme and a second substrate. In the case of NADH, an elegant approach is the application of formate dehydrogenase (FDH). This enzyme catalyzes the oxidation of formate anions to carbon dioxide with the concomitant reduction of NAD + to NADH. In such systems, NADH can be used for more than 80 000 cycles, although NADH costs are already negligible after 5000 cycles [22]. The advantage of FDH is the low cost of formate and the simple removal of the volatile reaction product, carbon dioxide, making the regeneration reaction virtually irreversible. Unfortunately, formate dehydrogenases are NAD + -dependent enzymes, but most P450s are dependent on NADPH. NADPH can be recycled, for example, with glucose dehydrogenase. Another approach is the use of an FDH mutant from Pseudomonas sp. 101, which was engineered to high activities towards NADP+ [23]. For further general information about the enzymatic cofactor regeneration see [6, 24]. Table 12.1 shows an overview of enzymatic cofactor regeneration systems that have been shown to drive P450 reactions. Taylor et al. immobilized a P450 enzyme from Streptomyces griseus (CYP105D1), expressed in E. coli, on the ionic exchange matrix DE52 and used an FDH as a recycling system for NADH regeneration [25]. The FDH was not immobilized and the model reaction was the hydroxylation of warfarin to 4-hydroxywarfarin. In the presence of the cofactor-regeneration system the amount of product was 5.5 times higher compared to the system where no recycling took place. This shows that the immobilized protein was active in bioconversions and that the requirement of a cofactor can be fulfilled by this recycling system for NADH regeneration. The immobilization of P450BM-3 from Bacillus megaterium in a

267

Regenerating enzyme

Formate dehydrogenase


Alcohol dehydrogenase

Glycerol dehydrogenase and alcohol dehydrogenase

P450

CYP105D1

P450BM-3

P450BM-3

P450cam

Ethanol

Acetaldehyde

Glycerol

GLD

NADH

NAP+

NADH P450

5-exo-Hydroxycamphor

Camphor

5-exo-Hydroxycamphor

Camphor

1,2-Epoxyhexane

1-Hexene

12-Oxo-dodecanoic acid + 4-Nitrophenol

p-Nitro-phenoxydodecanoic acid (surrogate substrate)

4-Hydroxywarfarin

Warfarin

P450

P450

P450

P450

NAP+

NADP+

NADPH

NADP+

NADPH

NAD+

NADH

ADH

ADH

FDH

FDH

Dihydroxyacetone

2-Propanol

Aceton

HCOO-

CO2

HCOO-

CO2

Reaction

Table 12.1 Overview of enzymatic P450 cofactor regeneration systems.

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25

Reference

268


Glucose-6-phosphate dehydrogenase

Glucose-6-phosphate dehydrogenase


Microsomal rat P450

P450BM-3

P450BM-3

G-6-P DH

HCOO-

CO2 FDH NADP+

NADPH

Arachidonic acid/ Linoleic acid

14(S),15(R)-Epoxyeicosatrienoic acid/ (+)-Leukotoxin

Glucose-6-phosphate

6-Phosphogluconate

P450

NADP+

NADPH

P450

Cyclohexanol

Cyclohexane

G-6-P DH

NADP+

NADPH

P450 7-Hydroxycoumarin

7-Ethoxycourmarin

7-Hydroxycoumarin

7-Ethoxycourmarin

32

31

30

12.3 Enzymatic Regeneration of Cofactors 269

270


sol–gel matrix was combined with a cofactor-recycling system based on NADP+ dependent formate dehydrogenase [26, 27]. The cytochrome was immobilized in a tetraethoxysilane (TEOS) sol–gel matrix and the formate dehydrogenase was separately immobilized in TEOS sol–gel matrix. The two enzymes were then mixed in a ratio of 1:1. This approach was used for the conversion of β-ionone, octane, and naphthalene to the respective hydroxy compounds with DMSO as cosolvent using P450BM-3 mutants. The sol–gel matrix-immobilized enzyme exhibited better storage stability. Recently the Arnold group described a biocatalytic system using P450BM-3 mutants to epoxidate 1-hexene to 1,2-epoxyhexane [28]. They used an alcohol dehydrogenase from Thermoanaerobium brockii for the cofactor recycling and 2propanol as cosubstrate. In this case the alcohol serves not only as the regeneration driving force, but also as the cosolvent for the hydrophobic substrate. A P450cam system with integrated enzymatic NADH regeneration was investigated in microscale aqueous compartments of stable water-in-oil emulsions formed by the non-ionic surfactant tetraethylene glycol dodecyl ether [29]. Two different dehydrogenases, bacterial glycerol dehydrogenase (GLD) and yeast alcohol dehydrogenase (ADH), were selected, and each of these was coupled with the P450cam catalytic cycle in water-in-oil emulsions. As a result, the camphor hydroxylation rate was successfully improved approximately 5-fold when GLD was employed under optimized conditions. These results reveal the potential utility of the microscale cell-like aqueous compartments of water-in-oil emulsions for multicomponent enzymatic reactions, especially for substrates with low solubility in water. Cytochrome P450 enzyme preparations obtained from phenobarbital-treated rats and glucose-6-phosphate dehydrogenase were co-immobilized in Romicon PM 10 polysulfone anisotropic hollow fibers [30]. The co-immobilization allowed cofactor recycling to take place but resulted in an apparent drop in the K m. The Vmax of immobilized preparations with or without cofactor recycling was about 25% of that observed with the non-immobilized preparations. Cytochrome P450BM-3 and its F87V mutant were exploited for preparation of 14(S),15(R)-epoxyeicosatrienoic acid [14(S),15(R)-EET] from arachidonic acid and (+)-leukotoxin B [(+)-12(S),13(R)-vernolic acid] from linoleic acid, respectively, by using glucose-6-phosphate dehydrogenase as the regenerating enzyme [31]. Maurer et al. have investigated the application of a mutant of P450BM-3 in a biphasic reaction system supported by cofactor recycling with NADP+ -dependent formate dehydrogenase from Pseudomonas sp. 101 [32]. Using this experimental set-up cyclohexane, octane, and myristic acid were hydroxylated. To reduce the process costs a novel NADH-dependent mutant of P450BM-3 was used in combination with an NAD + -dependent FDH. The stability of the monooxygenases under the reaction conditions is quite high, as revealed by the total turnover numbers of up to 12 850 in NADPH-dependent cyclohexane hydroxylation and up to 30 000 in NADH-dependent myristic acid oxidation.

12.4 Photochemical Approaches to Substituting or Regenerating Cofactors for P450 Systems

12.4 Photochemical Approaches to Substituting or Regenerating Cofactors for P450 Systems

Different photochemical approaches for cofactor regeneration and substitution have also been studied to achieve electron transfer to the cytochrome P450 systems. These processes were based on coupled light-active biomolecules such as chloroplasts, the photochemical reduction of flavins or mediated photo-induced electron transfer. A light-driven P450 monooxygenase system was prepared by mixing spinach chloroplasts and yeast microsomes containing a genetically engineered fusion enzyme between rat CYP1A1 and yeast P450 reductase [33]. Photosystem II in chloroplasts splits the water molecule, generating molecular oxygen, Photosystem I transfers electrons to NADP+ , yielding NADPH. The addition of ferredoxin (Fd) and ferredoxin-NADP reductase (FNR) was required to ensure that NADP+ was reduced to NADPH. The principle of the coupling system is shown in Fig. 12.1. A stable photoreaction system was also developed using cactus chloroplasts coupled with a fused P450/yeast reductase [34]. The immobilization of chloroplasts and yeast microsomes was tested to design a light-driven bioreactor [35]. Entrapment in agarose gave the best activity for the conversion of 7-ethoxycoumarin to 7-hydroxycoumarin under illumination of 6200 lx. The authors tested

Fig. 12.1 Principle of coupling P450 monooxygenases with chloroplasts as an artificial electron transfer system [34].

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three light-driven bioreactors; a two-phase column-type reactor with separately immobilized microsomes and chloroplasts showed the highest conversion rate compared with a reactor with co-immobilization of both components. The reactors showed turnover rates of 6.32 and 2.49 mol product mol−1 P450 min−1 after 40 and 180 min, respectively. Photochemical approaches to in situ regeneration of a cytochrome P450 monooxygenase via reduction of FAD have also been reported [36]. Riboflavin was covalently attached to the mammalian cytochrome CYP2B4, yielding an artificial flavocytochrome for photo-induced intermolecular electron transfer between the isoalloxazine cycle of flavins and the ferric heme group of cytochrome CYP2B4. The Gray group has succeeded in accelerating the delivery of electrons to the buried active site of cytochrome P450 by tethering a photochemical redox sensitizer, Ru-diimine, to a P450 substrate analog [37, 38]. In these Ru-diimine:P450 conjugates, the ruthenium donor and the ferriheme acceptor are held in position mainly by non-covalent interactions. Sevrioukova et al. achieved rapid photochemical heme reduction by covalently tethering a Ru-diimine to an engineered cysteine on the heme domain of P450BM-3 [39]. The authors did not investigate the biocatalysis of the reduced P450 in this system. A 6-fold higher electron transfer rate of the mediated photo-induced electron transfer from photoreduced tris(2,2′-bipyridyl)ruthenium(II) to the heme iron of engineered cytochrome CYP1A2 was measured compared with the wild type [40]. But even when electrons were transferred to the heme, no catalytic activity of 7-ethoxycoumarin de-ethylation was detected.

12.5 Electrochemical Systems for Substitution or Regeneration of Cofactors

Electrochemical enzyme studies are typically performed to determine fundamental parameters, such as the redox potential of the enzyme, or to study the electron transfer between an enzyme and an electrode. In addition, electrochemical studies of P450s are of great interest due to the possibility of developing applications such as biosensors and bioelectrochemical catalysts for product synthesis. Electrochemistry provides a powerful tool for electron transfer to P450 monooxygenases. Generally there are two ways in which a reaction is promoted by bioelectrocatalysis: direct electron transfer between electrodes and enzymes and indirect electron transfer using redox substances as mediators. Electrochemical approaches for electron transfer to P450 monooxygenases are often limited to anaerobic studies because oxygen is always likely to be a successful competitor for the electrons. In this case, only the fi rst one-electron reduction can be measured (Fig. 12.2). The direct electrochemistry of cytochrome P450 has been reported several times, but often without any proof of electrochemically driven substrate conversion [41–50]. Direct electrochemistry of cytochrome P450 and other heme enzymes on electrodes is generally difficult owing to the deeply buried heme center and instability of the enzyme upon interaction with the elec-


Fig. 12.2 Proposed electrochemically driven P450 catalytic cycle (adapted from [54]). RH, substrate; ROH, hydroxylated substrate. A – substrate binding, B – first reduction, C – O2 binding, D – second reduction, E – O2 – cleavage, F – product formation and release.

trode surface. A proposed electrocatalytic cycle of P450 is shown in Fig. 12.2. This mechanism requires that the electrode potential be sufficiently negative to reduce FeIII to FeII and to transfer the second electron to the Fe-oxygen after oxygen binding. For further literature on the general principles of the bioelectrochemistry of P450 monooxygenases see [10, 51–53]. 12.5.1 Electrochemical Regeneration of Natural Cofactors

Vilker and co-workers performed electrochemical regeneration of P450cam via cathodic reduction of putidaredoxin (Pdx) [55–57]. Pdx was chosen as a natural redox mediator on account of the difficulty of interacting directly with the interior heme of the large P450cam protein and in part because of the important role Pdx plays in maintaining the viability of the natural catalytic cycle (i.e. product stereochemistry, turnover rate, minimization of peroxide formation). Figure 12.3 shows the natural and direct electrode-driven catalytic cycle for camphor hydroxylation by the P450cam monooxygenase system. Vilker and colleagues used an antimony-doped tin oxide working electrode [56]. The required oxygen was produced at a platinum counterelectrode by water electrolysis in the anaerobic reaction system. In this way they minimized rapid reoxidation of putidaredoxin before it can deliver its reducing power to the P450. In addition, a tin oxide working electrode was screened by a platinum mesh to catalyze the decomposition of hydrogen peroxide which can diminish protein stability. A continuous catalytic cycle was sustained for more than 5 h and 2600 enzyme turnovers. The maximum product formation rate was 36 nmol of 5-exo-hydroxycamphor nmol−1 of P450cam min−1. This reaction concept was also applied to styrene epoxidation with a mutant of P450cam [55]. With a Pdx:P450cam ratio of

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Fig. 12.3 Natural and direct-electrode driven catalytic cycle for the P450cam monoxygenase system [56]. PdR, putidaredoxin reductase; Pdx, putidaredoxin; ox, oxidized; red, reduced; WE, working electrode; CE, counter electrode.

290:30 the reaction system produced 19 μmol L −1 styrene oxide after 4 h. At a ratio of 500:0.4 the concentration of styrene oxide produced was 30 μmol L −1 after 3 h. In this way the need for NADH and the flavin reductase was eliminated. Analogous experiments in the absence of Pdx showed that direct cathodic reduction of P450cam is possible but only at very slow rates [42]. Nevertheless the high excess of Pdx needed indicated that only a small portion of the coprotein in solution was actually involved in the catalytic reaction and a set-up with a higher electrode surface-to-reactor volume ratio would help to improve the bioelectrocatalytic efficiency. In mitochondrial P450, adrenodoxin (Adx) plays a similar role to putidaredoxin in the catalytic cycle. The direct unmediated electron transfer of Adx was shown at a gold electrode modified with a self-assembled layer of mercaptoundecanoic acid and with a polyelectrolyte layer of polyallylamine [58]. Direct electrochemistry of human adrenodoxin has been observed on a pyrolytic graphite electrode modified with poly-L-lysine [59]. This observation has not yet been used for the construction of a biocatalytic system. 12.5.2 Electrochemical Regeneration of Artificial Cofactors

Estabrook and co-workers showed that NADPH can be replaced by electrochemical regeneration of the mediator cobalt(II)sepulchrate for the electrocatalytically driven omega-hydroxylation of lauric acid [60, 61]. The reduced mediator can react with oxygen to form superoxide and hydrogen peroxide or reduce the cytochrome P450 complex. The hydrogen peroxide can be decomposed by the addition of catalase. Rates of formation of 12-hydroxydodecanoic acid by the electrochemical method are comparable to those obtained using NADPH as electron donor [62].


Incubation of the purified enzyme with the steroid progesterone in a reaction vessel containing a platinum electrode and an Ag/AgCl electrode couple poised at −650 mV, together with the redox mediator results in the 17-α-hydroxylation of the substrate at a rate of 25 nmol min−1 nmol−1 P450. A comparison of the rates of the reaction catalyzed by P450 using electrocatalysis or NADPH is given by [63]. For example, with P450BM-3, the substrate lauric acid, and NADPH as cofactor the hydroxylation rate was 900 nmol min−1 nmol−1 P450, whereas by using bioelectrocatalysis 110 nmol min−1 nmol−1 P450 was obtained. Applying different P450 and substrates, in each instance comparable reaction rates were achieved using NADPH or electrochemically supplied electrons. 12.5.3 Electrochemical Generation of Hydrogen Peroxide

An alternative transfer of the reduction equivalents from an electrode to the cytochrome P450 system was studied by reducing molecular oxygen to HO2− and H2O2 [64]. The electrochemically generated peroxide seems to be more active than added H2O2. In an electro-enzyme reactor several substrates were hydroxylated by microsomal cytochrome P450 with cathodically reduced oxygen which substitutes NADPH. Films of cytochrome CYP1A2 and polystyrene sulfonate were constructed on electrodes using layer-by-layer alternate absorption and evaluated for H2O2-driven enzyme-catalyzed oxidation of styrene to styrene oxide [16]. At −0.6 V vs. saturated calomel reference electrode (SCE) in an electrochemical cell, epoxidation of styrene was mediated by initial catalytic reduction of dioxygen to H2O2 which activates the enzyme for the catalytic oxidation. 12.5.4 Electrochemistry of P450 at Modified Electrodes

Electrochemical electron supply was used for the conversion of camphor to 5-exohydroxycamphor with P450cam immobilized at the electrode surface by means of a membrane [65]. The potential of the electrode was varied from 0.2 V to −0.8 V vs. SCE using a scan rate of 50 mV s−1. The formation of 5-exo-hydroxycamphor illustrated the feasibility of transferring electrons to monooxygenases in this way. In the presence of polylysine, spinach ferredoxin acts as an efficient electron shuttle between an indium-doped tin oxide (ITO) electrode and substrate-bound cytochrome P450cam [66]. Consequently, this strategy permitted the replacement of NADH and cytochrome P450 reductase with an electrode to catalyze the reductive dehalogenation of haloalkanes such as hexachloroethane. The positive electrostatic potential imparted by polylysine to the electrode surface assists spinach ferredoxin toward the electrode with an optimal orientation for heterogeneous electron exchange. Shumyantseva et al. reported that cytochromes CYP2B4 and CYP1A2 with a covalently attached riboflavin can be reduced electrochemically on rhodium–graphite electrodes at a potential of −500 mV vs. Ag/AgCl [67, 68]. N-

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Demethylation, p-hydroxylation, and O-dealkylation reactions proceeded in the presence of substrates such as aminopyrine, aniline, 7-ethoxyresorufin, and 7pentoxyresorufin, as was confirmed by product analysis. Rates of electrocatalytically driven reactions are comparable to those obtained using NAD(P)H as the source of reducing equivalents. The same group also investigated electron transfer between cytochrome CYP11A1 and gold nanoparticles immobilized on rhodium–graphite electrodes [69]. Thin fi lms of gold nanoparticles were deposited onto the rhodium–graphite electrodes by drop casting. Cytochrome CYP11A1 was deposited onto both gold nanoparticle-modified and bare rhodium–graphite electrodes. Cyclic voltammetry indicated enhanced activity of the enzyme at the gold nanoparticle-modified surface. On addition of aliquots of cholesterol to the electrolyte bioelectrocatalytic reduction currents were obtained. The gold nanoparticles enhanced electron transfer to the CYP11A1 when present on the rhodium–graphite electrodes. Using cyclovoltammetry, Lei et al. showed that P450cam, immobilized on a glassy carbon electrode modified with sodium montmorillonite, exhibits reversible redox behavior [44]. Thin films of sodium montmorillonite were also used with incorporated cytochrome CYP2B4, which were prepared on glassy carbon electrodes [70]. CYP2B4 can be reduced fast on clay-modified glassy carbon electrodes in the presence of Tween 80. In air-saturated solution, bioelectrocatalytic reduction currents can be obtained with the CYP2B4-modified electrode on addition of typical substrates such as aminopyrine and benzphetamine. The measurement of product formation indicates the successful bioelectrocatalysis by CYP2B4. The electrochemical behavior of human CYP2E1 absorbed or covalently linked to different electrode surfaces was investigated by Fantuzzi et al. [71]. Glassy-carbon and gold electrodes gave reversible electrochemical signals of an active CYP2E1. A monolayer coverage was obtained on gold modified with cystamine/maleimide that covalently linked surface accessible cysteines of CYP2E1. The turnover of the active enzyme was measured with the CYP2E1specific substrate p-nitrophenol, resulting in a K m of 130 μmol L −1 and the formation of 2.2 μmol L −1 of the p-nitrocatechol product upon application of a potential of −300 mV. Recently, the heme domain of P450BM-3 was electronically connected to a graphite electrode via a pyrene moiety covalently bound to the enzyme [72]. This method resulted in the fastest electron transfer rates observed so far for an electrode-bound P450 enzyme. 12.5.5 Electrochemistry of P450 in Surfactant Films

The majority of P450s are located in a hydrophobic environment in the cell. In order to mimic this physiological environment a number of groups have used phospholipids, such as didodecyl-dimethyl-ammonium-bromide (DDAB), for the construction of biosensors or bioreactors [9]. Phospholipid layers form stable ve-


sicular dispersions that bear a structural relationship to the components of the cellular membranes. By this means a membranous environment is created that facilitates electron transfer between the enzymes and the electrode. A biosensor containing cytochrome P450cam in a DDAB bromide vesicular system was prepared by cross-linking the enzyme onto a glassy carbon electrode with glutaraldehyde in the presence of bovine serum albumin [73]. Cyclic voltammetric responses of the sensor in an air-free buffer solution showed that the sensor exhibited reversible electrochemistry due to direct electron exchange between the heme FeIII/FeII redox system and the electrode surface. In air-saturated solution containing camphor, the biosensor gave an irreversible electrocatalytic current compatible with the monooxygenation of the substrate. The electrochemistry of human, bovine, and porcine cytochrome CYP17A1 has been examined on an edge-oriented pyrolytic graphite electrode [74]. The recombinant protein was immobilized on an electrode modified with DDAB. The P450 enzymes all retained electron-transfer activity while immobilized at the electrode surface as assessed by the presence of catalytic signals under aerobic conditions. Direct electrochemistry of P450 in a DDAB fi lm was also measured on a plasticformed carbon (PFC) electrode [75]. A quasi-reversible redox response was observed even at elevated temperatures of up to 80 ºC. The use of a thermophilic cytochrome P450, CYP119, in electrocatalytic dehalogenations of halocarbon solvents was studied. Temperature-stable enzyme-modified electrodes were constructed using sol–gel and polymeric surfactant approaches. CYP119 deposited in a dimethyldidodecylammonium poly(p-styrene sulfonate) (DDAPSS) fi lm has good retention of electrochemical activity up to 80 ºC [76]. At adequate potentials the CYP119/DDAPSS fi lms demonstrate high catalytic dehalogenation activities towards CCl4, CHCl3, and CH2Cl2. The Rusling group investigated protein–polyion fi lms grown layer-by-layer and cast protein–surfactant fi lms were employed on electrodes for catalytic oxidation of styrene derivatives to epoxides [15, 16, 77, 78]. P450cam in these fi lms mediated the electrochemical reduction of oxygen to hydrogen peroxide, which activates these heme proteins to catalyze olefin oxidation. Compared to bare electrodes with the proteins dissolved in solution, ultra-thin protein–polyion fi lms on gold electrodes coated with mercaptopropane sulfonate gave the best catalytic activities for the oxidations. Improved performance of protein–polyion fi lms is related to efficient, reversible heme FeIII/FeII electron transfer and better mechanical stability than the surfactant fi lms. Furthermore, dependence of product stereochemistry on oxygen availability in the reaction medium for the oxidation of cismethylstyrene suggested two pathways for olefin oxidation. The stereoselective pathway depends on an active, high-valent iron–oxygen intermediate as in the natural enzyme system, while the non-stereoselective pathway may involve a peroxyl radical near the protein surface. A composite amperometric biosensor was prepared by encapsulating a P450cam/DDAB liquid crystal system in a methyltriethoxysilane (MTEOS) sol– gel [79]. The composite sensing layer was cross-linked onto a glassy carbon electrode using glutaraldehyde. Analysis of the cyclic voltammetric results showed

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that the sensor exhibited faster reversible electrochemistry in the organic phase than in the aqueous phase. 12.5.6 Incorporation of Cytochrome P450 in Conducting Polymers

P450cam was immobilized without denaturing on an indium tin oxide electrode by polypyrrole and its activity was controlled electrochemically [80]. Polypyrrole, as a conductive polymer, was electropolymerized by passing a constant voltage of 3 V for 1 h in the presence of P450cam. Afterwards, when the potential of the electrode was swept repeatedly between 0.4 and 0 V vs. SCE, a decrease of oxygen in the reaction mixture solution was observed in the presence of camphor. In addition, hydroxycamphor was detected in the system by means of GC/MS. These results suggested that immobilized P450cam catalyzed the hydroxylation of camphor by the supply of electron from the electrode. An electrode system consisting of polyaniline-doped glassy carbon electrode and cytochrome CYP2D6 enzyme solution was used for the amperometric study of fluoxetine biotransformation [81]. The fi lm was grown potentiostatically at +700 mV vs. Ag/AgCl on a 0.071 cm2 glassy carbon electrode and used for cyclic voltammetric measurements. The polyaniline-mediated electrochemistry can be used to observe the monooxygenation reaction of CYP2D6. Summarizing this chapter, a large number of successful research approaches to electrochemically driven P450 catalysis have been reported to date, which can be taken as a sound “proof-of-principle”: the reductive dehalogenation of halogenated hydrocarbons [41, 66, 76], reduction of molecular oxygen [45, 46, 64, 72], reduction of nitric oxide [82], epoxidation of styrene [15, 16, 55, 77, 78], hydroxylation of fatty acids [60, 62, 63], hydroxylation of steroids [60, 63, 74], hydroxylation of nitrophenol [71] and hydroxylation of camphor [56, 65, 73, 80] and also the demethylation and dealkylation of drugs [60, 67, 68, 81].

12.6 Redox Mediators

Redox mediators can be used for the efficient electron transfer from an electrode, a photochemical system or an artificial electron donator to P450. In each case the mediator must be able to be reduced (e.g. at an electrode at an appropriate potential), and the reduced form must be able to transfer the electrons to an enzyme. A frequently used mediator for oxygenation reactions catalyzed by P450 is cobalt(II)sepulchrate (Fig. 12.3). The organometallic one-electron transfer reagent has been regenerated either electrochemically [60–62, 73] or by chemical reductants such as elementary metals [83, 84] and coupled to P450 for various hydroxylation reactions. In each of these approaches the monooxygenase was used together with a reductase either as a fusion protein or with a soluble reductase in a reconstituted bienzymatic reaction system. Thus, the pathway of electron flow in these

12.6 Redox Mediators

examples does not necessarily include the direct reduction of the heme prosthetic group but more likely proceeds via reduction of the flavin-dependent reductase, which delivers the monooxygenating subunit with the reducing equivalents needed for reductive activation of O2 [62]. The feasibility of replacing NADPH with 1,1′-dicarboxycobaltocene in the catalytic cycle of P450BM-3 has been explored [85]. Using the holoprotein, the mediator was shown to reduce both the FAD and FMN in the reductase domain as well as the iron in the heme domain. In an electrochemical system, the mediator was able to support lauric acid hydroxylation at a rate of 16.5 nmol product nmol−1 enzyme min−1. Similar electron transfer and catalysis were observed for the heme domain alone in the presence of the metallocene. Parallel studies under the same conditions using a cobalt sepulchrate mediator showed that the two systems give similar results for both the holoenzyme and the heme domain. Tris-2,2-bipyridine ruthenium(II) (Ru(bpy)32+ ) was frequently used as photo-reduced mediator for P450 [39, 40, 86]. The catalytic cycle of wild-type P450cam can be supported in the presence of the artificial reductant potassium ferrocyanide [87]. Further mediators for P450 are neutral red [88], methylviologen [89], and phenosafranine [90]. Figure 12.4 shows the corresponding chemical structures.

Fig. 12.4 Structures of different mediators used as electron donators for P450 monooxygenases.

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Further development of the electron transfer system can be expected by using directed evolution methods to improve the P450 performance towards mediated electron transfer [91]. A validated medium-throughput screening system was designed in order to improve P450 monooxygenases by mediated electron transfer with zinc/cobalt(III)sepulchrate as an alternative and cost-effective cofactor system. The mutant P450BM-3 F87A was used as a model system for developing the screening system in a 96-well format. The mediator evolution screen was validated by comparing the activity of P450BM-3 to P450BM-3 F87A and by screening a saturation mutagenesis library at amino acid position R47. For mediated electron transfer, two double mutants, P450BM-3 (F87A R47F) and P450BM-3 (F87A R47Y), were identified with 2- to 3-fold increased catalytic efficiency. Mediators can also be used for the electrochemical regeneration of cofactors [92–94]. Nevertheless, as the direct reduction of nicotinamide cofactors usually leads to inactive dimers and byproducts [95], indirect methods (using e.g. rhodium complexes as mediators) are more promising.

12.7 Molecular Biological Approaches 12.7.1 Peroxide Shunt

The directed evolution of monooxygenases offers the possibility of optimizing the enzymes towards a more efficient peroxide shunt pathway and a higher enzyme stability. The replacement of phenylalanine 87 with alanine or glycine increased the peroxide-supported substrate hydroxylation of P450BM-3 [96, 97]. Using the enzyme wild type (WT) the turnover number of the peroxide-driven hydroxylation of the surrogate substrate pNCA (p-nitrophenoxydodecanoic acid) is lower than 0.5 nmol min−1 nmol−1 P450 [96]. The mutants F87A and F87G show a turnover number of 162 ± 11 and 78 ± 9 nmol min−1 nmol−1 enzyme, respectively. Compared with the natural cofactor NADPH the activities of F87A and F87G were 37 and 43%, respectively. The inactivation of F87A and WT in the substrate-free form and the substrate binding complexes has been investigated with various concentrations of hydrogen peroxide by analyzing the remaining activity. Substrate-free WT in 1 mmol L −1 H2O2 lost about 55% of its activity within 7 min, while F87A in 16 mmol L −1 only lost 45% during the same period. The inactivation of the F87A substrate-binding complex was much slower than that of the WT substrate-binding complex in H2O2 solutions. Similarly the substrate-free F87A was more stable than substrate-free WT in the same solutions. The speeds of inactivation of the WT or F87A substrate-binding complexes were significantly faster than their respective substratefree controls. Arnold and co-workers have succeeded in the evolution of P450cam and P450BM-3 with drastically improved activity and stability towards peroxide

12.7 Molecular Biological Approaches

[97–99]. A mutant of P450cam shows a 20-fold higher hydroxylation rate of naphthalene than the wild-type enzyme [98]. The P450BM-3 heme domain (BMP) supports peroxygenase activity and is more thermostable than the whole P450BM-3. The usage of the BMP mutant F87A as the starting point to increase catalyst performance by sequential rounds of random mutagenesis and screening for hydrogen peroxide-driven hydroxylation of pNCA led to an approximately 20-fold higher activity in 10 mmol L −1 H2O2 [99]. 12.7.2 Artificial Electron Transfer Systems

An artificial electron transfer system has been constructed from thermostable CYP119 of the extremophilic archaebacteria Sulfolobus sulfataricus, putidaredoxin, and putidaredoxin reductase [100]. Neither the natural redox partners of CYP119 in Sulfolobus sulfataricus nor the endogenous substrates are known. In the absence of this information, the catalytic activity of the cytochrome has been measured using styrene as the substrate in conjunction with putidaredoxin and putidaredoxin reductase, the electron transfer partners of P450cam. Gilardi and coworkers assembled an artificial redox chain in a chimeric fusion protein, using the heme domain of CYP102 and flavodoxin from Desulfovibrio vulgaris connected by a natural loop of CYP102 reductase; the working electrode was a glassy carbon disc with platinum wire [101–104]. The idea was to assemble non-physiological redox modules, generating artificial redox chains, which could lead to new multidomain constructs with improved electrochemical and catalytic properties. Semisynthetic flavocytochromes, likewise desribed above [67], obtained by covalent binding of riboflavin with cytochromes CYP2B4, were able to catalyze the H2O2mediated reactions of aniline, aminopyrine and p-nitroanizole [17]. 12.7.3 Changing the Cofactor Specificity of P450 Systems

Another way to overcome the high cost for NADPH is to change the specificity of P450 to NADH. NADH as cofactor is about 10 times cheaper than NADPH, more stable, and furthermore can be recycled by a wider variety of dehydrogenases. P450BM-3 shows a strong preference for NADPH over NADH as reducing coenzyme. An aromatic amino acid is located over the FAD isoalloxazine ring, and in some cases it is important in controlling coenzyme specificity [105]. In P450BM-3 the aromatic residue inferred from sequence alignments to stack over the FAD is Trp1046; mutation to Ala1046 and His1046 effected a remarkable coenzyme specificity switch. Eiben et al. have investigated different single and double mutations at the NADPH-binding domain [106]. To identify NADH-dependent mutants, kinetic constants for the reduction of cytochrome c with NADPH and NADH were determined. Catalytic efficiency with NADH was improved 600-fold compared with the starting point. One mutant was used in an enzyme reactor for the conversion

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of dodecanoic acid. NADH was also recycled with NAD + -dependent formate dehydrogenase. With a total turnover number of almost 70 000, a space–time yield of 3.3 g hydroxylated laurate per litre in 24 h was achieved. A functional human NADH-dependent cytochrome P450 system has been developed by altering the cofactor preference of human NADPH cytochrome P450 reductase [107]. This has been achieved by a single amino acid change in the conserved aromatic amino acid Trp676, which covers the FAD isoalloxazine ring in the nicotinamide-binding site. Of the mutations made, the substitution of Trp676 with alanine (W676A) resulted in a functional NADH-dependent enzyme, which catalyzed the reduction of cytochrome c and ferricyanide as well as facilitated the metabolism of 7-ethoxyresorufin by CYP1A2. Kinetic analysis measuring cytochrome c activity revealed that the NADH-dependent kcat of W676A is equivalent (90%) to the NADPHdependent kcat of the wild-type enzyme, with W676A having an approximately 1000-fold higher specificity for NADH. 12.7.4 Intracellular Cofactor Regeneration

Recently metabolic engineering has been used to improve the catalytic performance of a P450 whole cell biocatalyst by introducing a heterologous NADH regeneration system, glycerol dehydrogenase (GLD) [108]. Recombinant E. coli harboring the P450cam system, but lacking GLD, exhibited little activity for camphor hydroxylation. In contrast, coexpression of GLD resulted in an almost 10-fold improvement of substrate conversion, implying that the whole cell biocatalyst utilized molecular oxygen, endogenous NADH, and glycerol for catalysis. The addition of glycerol to the reaction media further promoted camphor hydroxylation, suggesting that exogenous glycerol is also available for GLD in the host cell and actively participates in the catalytic cycle.

12.8 Conclusion and Outlook

At present, industrial processes with P450 monooxygenases (e.g. steroid biotransformations) are dominated by whole cell biocatalysis [109, 110] due to the technical difficulties which arise from the use of isolated P450 (i.e. enzyme stability and cofactor (often NADPH) dependence), and the obvious advantages of intact cell systems in this context. However, there are also significant drawbacks associated with whole cell P450 processes (e.g. limited cellular uptake and export of hydrophobic substances, byproduct formation by metabolic side reactions and product degradation, cytotoxicity of hydrophobic substrates and products due to membrane damage). The development of different systems for the regeneration and substitution of the natural P450 cofactors may help granting access to technical in vitro P450 applications in the future. Table 12.2 summarizes the advantages and disadvantages

12.8 Conclusion and Outlook

283

Table 12.2 Advantages and disadvantages of various strategies for cofactor regeneration and substitution.

Strategy

Advantages

Disadvantages

Enzymatic regeneration

High selectivity Compatibility to P450 reaction conditions High turnover rates possible Low cost of electricity

Enzyme cost and instability Higher complexity of reaction system

Electrochemical regeneration

Photochemical regeneration

Substitution by peroxides

No stoichiometric regenerating reagent Easy product isolation Easy monitoring of reaction progress Inexpensive No stoichiometric regenerating reagent in some systems

Low price for peroxides Simple construction of reactions systems

Direct electrochemistry

Low cost of electricity No stoichiometric regenerating reagent Easy product isolation Easy monitoring of reaction progress

Mediated electron transfer

No further chemicals or mediators necessary Fast electron transfer Wide range of mediators Facile interaction with buried active sites

Product purification may be complicated Incompatibility with some biochemical systems Poor selectivity (especially for reductive regeneration) Complex apparatus and procedures Fouling of electrode Complex apparatus Limited compatibility with biochemical systems Requirement for photosensitizers and redox dyes Low stability of the P450 towards hydrogen peroxide Strong oxidizing agent capable of reacting directly and non-enzymatically with substrates Active site deeply buried in protein Dependence of electron transfer on protein orientation on the electrode surface Adsorptive denaturation on metal electrodes Formation of an insulating protein layer which prevents electron exchange with electrodes

Mediator cost Must be immobilized or removed to avoid product contamination Regeneration system for the mediator needed

Partially adapted after [10, 112].

of the different major strategies discussed in this paper. It is not possible to give a general recommendation for the best strategy since each reaction system has to be evaluated based on its characteristic demands (type of P450 needed, substrate and product properties, purification requirements, etc.). Clearly, enzymatic cofac-

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tor regeneration systems are of high industrial potential and have already proven to be well suited for commercial applications, as is the case, for instance, with NADH-dependent alcohol dehydrogenase-catalyzed syntheses coupled to formate dehydrogenase (FDH) or glucose dehydrogenase [111]. Enzymatic NADPH regeneration systems are less developed, and promising approaches (e.g. the use of engineered NADP+ -dependent FDH) have been recently reported [23]. Nevertheless, besides major tasks associated with the development of in vitro P450 catalysis, especially those concerning activity, specificity, and stability, the enzyme’s demand for the expensive natural cofactor NAD(P)H is probably the most critical issue from the economist’s viewpoint. Thus, it seems worth investigating technical solutions to achieve the ultimate goal, the complete elimination of the cofactor from the reaction system. Bioelectrochemistry can certainly provide clean and cheap solutions for in vitro P450 catalysis but this demands further joint research efforts at the interface of the disciplines electrochemistry and molecular biology/biocatalysis. The current state-of-the-art of electrochemically driven P450 catalysis can be taken as “proofof-principle” that NAD(P)H dependence of P450 can be overcome by “electrons from the wall socket.” Nevertheless, extensive research activities are still needed to pave the way towards more application-oriented developments. Here, among many technical issues, such as electrode-to-enzyme electron transfer, operational stability of P450, etc., it is the proper design of an electroenzymatic reactor allowing for space–time yields comparable to those known from conventional biocatalytic systems, which may be the most critical parameter. Together with the steadily increasing knowledge of the molecular mechanisms of P450 catalysis and the sophisticated enzyme engineering tools, such as directed evolution and rational design, even smarter cofactor substitution systems can be expected in the future and may eventually help P450 monooxygenases perform “modern biooxidations” as in vitro biocatalysts on an industrial scale. References 1 Hollmann, F., Hofstetter, K., Schmid, A. Non-enzymatic regeneration of nicotinamide and flavin cofactors for monooxygenase catalysis. Trends Biotechnol 2006, 24, 163–171. 2 Zhao, H., van der Donk, W. Regeneration of cofactors for use in biocatalysis. Curr Opin Biotechnol 2003, 14, 583–589. 3 Hanukoglu, I. Electron transfer proteins of cytochrome P450 systems. Adv Mol Cell Biol 1996, 14, 29–55. 4 van der Donk, W., Zhao, H. Recent developments in

pyridine nucleotide regeneration. Curr Opin Biotechnol 2003, 14, 421–426. 5 Adlercreutz, P. Cofactor regeneration in biocatalysis in organic media. Biocatal Biotrans 1996, 14, 1–30. 6 Eckstein, M., Daussmann, T., Kragl, U. Recent developments in NAD(P)H regeneration for enzymatic reductions in one- and two-phase systems. Biocatal Biotrans 2004, 22, 89–96. 7 Chenault, H., Simon, E., Whitesides, G. Cofactor regeneration for enzymecatalysed synthesis. Biotechnol Genet Eng Rev 1988, 6, 221–270.

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291

Index

a abortive P450 catalytic cycles 142 acetaldehyde 9 acetate peroxidation 8 acetic acid bacteria – biooxidation 1 – D -mannitol dehydrogenase 21 – D -sorbitol dehydrogenase 22 Acetobacter – alcohol oxidation 7 – aldehyde dehydrogenase 33 acetylcholinesterase 63 Achromobacter xylosoxidans 202 Acinetobacter – alcohol dehydrogenase 10 – Baeyer–Villiger monooxygenases 80 – D -hexosamine dehydrogenase 30 Acinetobacter calcoaceticus – glucose oxidation 11 – quinate oxidation 24 – s-GDH 12 actinomycetes 179 ADH, see alcohol dehydrogenase adrenodoxin (Adx) 128 adrenodoxin reductase (AdR) 127–128, 163 aerobic fermentation system 197 aflatoxin biosynthesis 92 alcohol dehydrogenases (ADH) 9 – Acinetobacter 10 – cyclic 9 – membrane-bound 6 – quinohemoprotein 7 alcohol oxidation 6 – acetaldehyde 9 – Acetobacter 7 – aldehyde 9

– aliphatic ketone 9 – aliphatic secondary alcohol 9 – cyclic ketone 9 – direct electron transfer 8 – ethanol 9 – Gluconobacter 7 – membrane-bound alcohol dehydrogenase 9 – quinohemoprotein 6 – secondary alcohol dehydrogenase 9 alcohol sensors, ADH III 8 aldehyde, alcohol oxidation 9 aldehyde dehydrogenase (ALDH) 3 – Gluconobacter 34 – membrane-bound 33 alicyclics 109 aliphatic dicarboxylic acids, industrial oxidation 196 aliphatic ketone 9 aliphatic secondary alcohol 9 alkanes 197 amino acid alignment 134 amperometric biosensors, electrochemistry 277 androstane – basic structure 156 – biotransformations 155 anthrax 62 application modes, laccase 53 Arabidopsis thaliana 170 D -arabitol 14 arabitol 1 dehydrogenase (ARDH) 14 aromatic compounds – 3,4-dimethyl-benzaldehyde 200 – catechols 198–199 – conversion of toxic compounds 198–199 – heterocyclic 201

292

Index – (R)-2-(4-hydroxyphenoxy)propionic acid (HPOPS) 199 – selective oxidation 200 – styrene oxide production 200 – two-liquid phase system 200 Arthrobacter oxydans 202 artificial cofactors 274–275 artificial electron transfer systems 281 L-ascorbate 23 Aspergillus fumigatus, 11α-hydroxylation 161 Aspergillus niger, 11α-hydroxylation 160 Aspergillus parasiticus 92 atomic structures – eukaryotic nitric oxide synthase 140 – nitric oxide synthase 140 – P450 (heme) domain 132 – P450BM-3 104 – P450cam 127 – rat CPR 129

b Bacillus megaterium 158 – P450 redox partners 125, 131 – P450BM-3 134 – steroid biotransformation 164 Bacillus subtilis, P450BM-3 134 bacterial laccases 59 bacterial P450 monooxygenases 99 – expression 180–181, 183–186 – functional diversity 178 – reaction array 186–187 Baeyer–Villiger monooxygenases (BVMO) 77–94 – genome mining 92 – PAMO 82 – peracid catalyst 79 – peroxyflavin enzyme intermediate 81 – see also type I BVMOs Baeyer–Villiger reaction 77 batch fermentations 197 batteries 64 Beauveria bassiana 228 Beauveria bassica 199 Bertrand–Hudson’s rule, polyol oxidation 15 biological warfare agents 62 bioreactors 233–252 – drug discovery 239–250 – stainless steel 248 biorefinery 66 bioremediation 50

biosensors – amperometric 277 – Erwinia sp. 13 – laccase 53 – see also sensors bleaching 50, 56 blood D -glucose monitoring 13 BVMO, see Baeyer–Villiger monooxygenases

c camphor 102 Candida albicans 218 Candida antarctica 79 Candida tropicalis 195 – dicarboxylic acids 197 – industrial oxidation 196 carbomycin A 181 carbon electrode, conducting polymers 278 carbon source addition, Roche experience 242 carboxylic acid 33 cardboard packaging materials 55 cardboard strengthening 56 catalysed oxidations, drug metabolism 219–220 catalysts, whole cells 86 catalytic efficiency, P450 redox partners 144 catechols 198–199 cationic species 65 CBZ (carbamazipine), epoxidation 226 charge-pair interaction 159 chemical warfare agents 62 chlorzoxazone, Roche experience 242 CHMO, see cyclohexanone monooxygenase clustering 84 Cochliobolus lunatus, 11β-hydroxylation 161 cofactor regeneration 268 – electrochemical 273–275 – enzymatic 271 – photochemical 271–272 cofactors – costs 266 – flavin 78 – NAD(P)H 267–271 – non-covalently bound 144 – specificity change 281 – substitution 266–271 colorant bleaching 50 conducting polymers 278 cork stoppers 56 coupled reaction 267

Index coupling efficiency 105 coupling reaction 16 CPR, see CYP reductase Criegee intermediate 77 crop protection 51 Cunninghamella blakesleeana, 11βhydroxylation 161 Cunninghamella elegans, microbial hydroxylations 228 Curvularia lunata, 11β-hydroxylation 161 cyclic alcohols 9–10 cyclic ketone 9 cyclohexanone monooxygenase (CHMO), BVMO 80 cyclovoltammetry 276 CYP, see P450 (cytochrome P450 monooxigenase) – see also Baeyer–Villiger monooxygenases CYP engineering database (CYPED) 254–257 CYP reaction array – biotransformation 182–183, 187–190 – construction 186–187 CYP reaction screening 224 CYP101, see P450cam CYP102 (A1, A2, A3), see P450BM-3 CYP105 189 CYP105D1 267 CYP106A2 165 CYP107C1 183 CYP107C1 expression 181 CYP107H1 137 CYP116B2 136 CYP11B 167 CYP1A2 221 CYP2D6 alleles 257 CYP3A4 246–247 CYPED, see CYP engineering database cytochrome P450 monooxygenase, see P450 cytosolic fractions, biooxidation 4

d DAHP 228 Data Warehouse ARchitecture for protein classiFication (DWARF) 254 DCA, see dicarboxylic acids DCIP, see dichlorophenol indophenol decontamination systems 62 dehydrogenases – biooxidation 1 – FAD-dependent 27 – membrane-bound 2

– polyol oxidation 23 – PQQ-dependent 6 3-dehydroquinate 24 delignification 49 denim bleaching 56 N -desethyl amodiaquine 242 development costs 57 dextromethorphan 259 DHA, see dihydroxyacetone diabetes 13 dicarboxylic acids (DCA) 197–198 – aerobic fermentation system 197 – alkanes 197–198 – batch fermentations 197 – Candida tropicalis 197 – industrial oxidation 196–198 – β-oxidation pathway 196 dichlorophenol indophenol (DCIP) 5 10,11-dihydro-10-hydroxy-carbamazepine 225 dihydroxyacetone – industrial oxidation 206 – polyol oxidation 19 3,4-dimethyl-benzaldehyde 200 2,5-dimethylpyrazine 201 direct electron transfer 8 direct-electrode driven catalytic cycle 273–274 directed evolution 66, 83, 100, 116, 165, 191 dithionite, NAD(P)H 266 DMEs, see drug-metabolizing enzymes drug development 233–252 drug discovery – bioreactors 239–250 – Escherichia coli 235, 238–250 – enzymic activities 237–239 – episomally replicating vectors 236 – expression 239–250 – P450 expression systems 239–250 – preclinical development 234 – Roche experience 239 – Saccharomyces cerevisiae 238 drug metabolism – CYP isozyme-specific reactions 221–225 – eukaryotic P450s 217–221 – human 234 – human DME system reaction 217 – microbial hydroxylations 225–228 – microbial vs. mamalian 221–225 – reaction screening 224 drug metabolites 211–232 drug-metabolizing enzymes (DMEs) 211–213

293

294

Index drug preparation 211 DWARF, see Data Warehouse ARchitecture for protein classiFication dye bleaching 50 dye precursors 52

e E. coli, see Escherichia coli electrocatalysis 141 electrochemical generation, see cofactor regeneration electrochemical systems 272–278 electrochemistry 275–278 – amperometric biosensors 277 – glassy carbon electrodes 276 electron donators – P450 redox partners 137 – redox mediators 279 electron transfer – glucose oxidation 12 – NAD(P)H 281 – P450 redox partners 125 engineering, laccase 66 environmental contaminants, P450 107 enzymatic fuel cells 64 enzyme assay, laccase 53 enzyme cooperation, laccase 62 episomally replicating vectors 236 epoxidation 226 10,11-epoxy-carbamazepine 225 error-prone PCR 116 Erwinia sp. 13–14 meso-erythritol oxidation dehydrogenase – Gluconobacter frateurii 17 – membrane-bound 16 L-erythrulose 16 Escherichia coli, bacterial CYP expression system 183–185 – biotransformation 177, 185 – expression library 185 – Novartis experience 244 – recombinant 240 ethanol oxidation, see alcohol oxidation ethionamide monooxygenase 86 eukaryotic microorganisms 216

g

f FAD-dependent dehydrogenase 27 – biooxidation 1 – formaldehyde-oxidizing enzyme – 5-keto- D -fructose 27 – D -galactosaminate 29 – D -galactosamine 29

– 2-keto- D -gluconate 28 – D -glucosaminate 29 – D -glucosamine 29 – D -mannosaminate 29 – D -mannosamine 29 – membrane-bound 28, 32 – sorbitol 32 fatty acids – industrial oxidation 194–198 FDH, see formate dehydrogenase fermentation – biooxidation 1 – effluent 195 – industrial oxidation 195 – laccase 67 – Roche experience 240 ferredoxins 137–138 fi lms 276–278 fine chemicals 198–200 flavin adenine dinucleotide, see FAD flavodoxins 137–138 flavoenzymes 66 flavoprotein 78 formaldehyde-oxidizing enzyme 34 formate dehydrogenase (FDH) 271 fossil fuel pollutants 63 D -fructose 20 5-keto- D -fructose 27 D -fructose dehydrogenase 27 fungal laccases 43, 48 fungal strains 211–232 fusion enzymes – biocatalytic potential 138–140 – CYP116 132 – novel class 136–137 – P450BM-3 102

34

GADH, see D -gluconate dehydrogenase D -galactosaminate 29 D -galactosamine 29 GDH, see D -glucose dehydrogenase m-GDH, oxidation 11 s-GDH – Acinetobacter calcoaceticus 12 – oxidation 12 genetically modified microorganisms 163–170 genome mining, BVMOs 92 glassy carbon electrodes – conducting polymers 278 – electrochemistry 276 GLDH, see glycerol dehydrogenase

Index D -gluconate 12 2,5-diketo- D -gluconate 31 2-keto- D -gluconate, FAD-dependent dehydrogenase 28 5-keto- D -gluconate, polyol oxidation 17 D -gluconate accumulation 18 D -gluconate dehydrogenase (GADH) 31 – activation 29 – membrane-bound 28 2-keto- D -gluconate dehydrogenase 31 D -gluconate oxidation 16 D -gluconate oxidizing polyol dehydrogenase 17 Gluconobacter – alcohol oxidation 7 – aldehyde dehydrogenase 34 – glucose oxidation 11 – polyol oxidation 17 Gluconobacter frateurii 10 – meso-erythritol oxidation dehydrogenase 17 – D -hexosamine dehydrogenase 31 – polyol oxidation 21 Gluconobacter industrius – GLDH 19 – polyol oxidation 15 Gluconobacter melanogenus 31 Gluconobacter oxydans – glycerol conversion 206 – polyol oxidation 23 – quinate oxidation 24 Gluconobacter suboxydans, polyol oxidation 14 D -glucono-δ-lactone, oxidation 11–12 D -glucosaminate 29 D -glucosamine 29 D -glucose – oxidation 11–12 – sensors 13 D -glucose dehydrogenase (GDH) – sensors 13 – soluble 12 glucose oxidation 11 – Acinetobacter calcoaceticus 11 – D -arabitol 14 – blood D -glucose monitoring 13 – electron transfer 12 – D -glucono-δ-lactone 11–12 – D -glucose 11–12 – m-GDH 11 – Pseudomonas 11 – quinoprotein GDHs 13 – s-GDH 12

– sensors 13 – D -xylulose 14 glycerol conversion 206 glycerol dehydrogenase (GLDH) – enzyme purification 20 – Gluconobacter industrius 19 – polyol oxidation 18 gonane, structure 155

h HAPMO 83 heme absorption band 124–125 heme B-containing monooxygenase, CYP 123 heme iron spin-state shift 125 heterologous expression – bacteria 235 – yeast 236–237 D -hexosamine dehydrogenase – Acinetobacter 30 – G. frateurii 31 – membrane-bound 29 hormonal steroids, biotransformations 155 HPOPS, see (R )-2-(4hydroxyphenoxy)propionic acid HQ, see hydroquinone human CYP 215 – metabolites 114 – model 253–264 human DME system reaction 217 human drug metabolism 234 human P450 expression 233–252 hydrocarbons 194–198 hydrogen peroxide – electrochemical generation 275 hydroquinone (HQ) state 137 4,5-bis-(4′-hydroxyanilino)-phthalimide 227–228 7-hydroxycoumarin 182 11α-hydroxylation 160–162 – Aspergillus fumigatus 161 – Aspergillus niger 160 11β-hydroxylation – Cochliobolus lunatus 161 – Curvularia lunata 161 16α-hydroxylation 162 6-hydroxy-(S )-nicotine, heterocyclic compounds 202–203 6-hydroxynicotinic acid 202 (R )-2-(4-hydroxyphenoxy)propionic acid (HPOPS) 199 5-hydroxypyrazinecarboxylic acid 202 4-[6-hydroxypyridin-3-yl]4-oxobutyrate 202–203

295

296

Index

i indigo 203 indole 204 industrial oxidation processes – alcohols 194 – aliphatic dicarboxylic acids 196 – alkanes 194–198 – aromatic compounds 198–200 – Candida tropicalis 196 – dicarboxylic acids 196–198 – dihydroxyacetone 206 – fatty acids 194–198 – fermentation processes 195 – fine chemicals 198–200 – glycerol conversion 206 – heterocyclic compounds 200 – hydrocarbons 194–198 – β-oxidation pathway 194 – yeasts 194 intermediates – Criegee 77 – peroxyflavin enzyme 81 – rearrangement 78 intracellular cofactor regeneration 282

k Ketogluconicigenium vulgare

23

l laccase 43 – application modes 53 – engineering 66 – enzyme assay 53 – enzyme cooperation 62 – fermentation 67 – mediated laccase oxidation 64 – mediators 54 – Myceliophthora thermophila 44 – Nephotettix cincticeps 58 – Novozyme 51003 56 – Panus tigrinus 57 – Pfam database 58 – Pleurotus ostreatus 57 – protein engineering 66 – radical species 65 – structure 43 – TEMPO 60, 65 – Trametes villosa 45 – Tricholoma giganteum 58 – white laccase 57 – yellow laccase 57 large-scale pharmaceutical biooxidations 233

lignin, laccase 49 link sequence, computational analysis 58 lipids degradation, laccase 63 long-chain aliphatic dicarboxylic acids 196

m D -mannitol

dehydrogenase 20–21 29 D -mannosamine 29 mediators, laccase 54 medium-chain alcohols, industrial oxidation 194 membrane fractions – biooxidation 4 – EDTA treatment 5 membrane-bound – alcohol dehydrogenase 6, 9 – aldehyde dehydrogenase 33 – arabitol 1 dehydrogenase 14 – dehydrogenases 2 – meso-erythritol oxidation dehydrogenase 16 – D -fructose dehydrogenase 27 – GLDH 15, 19 – D -gluconate dehydrogenase 28 – 2-keto- D -gluconate dehydrogenase 31 – D -gluconate oxidizing polyol dehydrogenase 17 – D -glucose dehydrogenase 11 – D -hexosamine dehydrogenase 29 – D -mannitol dehydrogenase 20 – P450 166–170 – quinate dehydrogenase 24 – ribitol dehydrogenase 21 – sorbitol dehydrogenase 22, 32 – L-sorbosone dehydrogenase 23 milbemycins 229–230 mitochondria 128 mitochondrial steroid hydroxylases 159 modified electrodes 275 monooxygenases – Baeyer–Villiger 77–94 – CYP, see P450 MtmOIV, BVMO 79 multi-copper oxidases 43 Myceliophthora thermophila 44 Mycobacterium tuberculosis 92 mycotoxins 90 D -mannosaminate

n NAD(P)H 87, 99, 125, 161 – dependent enzymes 4

Index – electron transfer systems 281 – enzymatic regeneration 267–271 – recycling 265–290 naphthalene oxidation 108 Neotyphodium uncinatum 91 Nephotettix cincticeps 58 niacin, heterocyclic compounds 202 nitric oxide synthase (NOS) enzymes 139 Nonomuraea recticatena 177 NOS, see nitric oxide synthase Novartis 217, 248 Novozyme 51003 56

o oleanolic acid 189 organophosphorus compounds oseltamivir 25 overoxidation 88 β-oxidation pathway – dicarboxylic acids 196 oxidizing agent 77

63

p P420 P450

144 99, 123, 143, 215 – alicyclics 109 – alkanes 109 – aromatic compounds 105 – Bacillus megaterium 99 – biocatalysis 88 – biodiversity 123, 143 – biotechnology 123 – biotransformations 155–176 – camphor 102 – classes 158 – cofactors, see cofactors – conducting polymers 278 – coupling efficiency 105 – cycle 125 – directed evolution 116 – engineering 116 – environmental contaminants 107 – enzymatic regeneration 100 – enzyme engineering 109 – functional expression 101 – genes 178 – heme B-containing monooxygenase 123 – human metabolites 114 – inactive P420 form 144 – industrially valuable 179 – membrane-bound 166–170 – microbial 177–192

– models 253–264 – monoterpene binding 111 – mutation 106 – nomenclature 253 – novel types 132 – pharma research 239 – photochemical approaches 271–272 – polychlorinated aromatics 107 – protein components 124 – protein engineering 130 – Pseudomonas putida 99 – redox partners 123 – selectivity improvement 113 – stability 144 – steroid hydroxylase systems 156–159 – terpenoid compounds 111 – utility 239 – Y96A mutation 106 P450 (heme) domain 132 P450 monooxygenases, see P450 P450 mutants, active site structures 108 P450 redox partners 123–124 – catalytic cycle 125–127 – coenzymes 142–143 – CYP116B2 136 – electron donor system 137 – electron transfer events 125 – enzyme stability 142–143 – ferredoxins 137–138 – flavodoxins 137–138 – fusion enzymes 131, 136–137, 138–140 – stability 142–143 P450BM-3 99, 131, 158 – active site structure 104 – amino acid alignment 134 – Bacillus megaterium 134 – biocatalytic potential 138 – general features 102 – mutagenesis studies 115, 131, 271–272 – physiological role 133 P450cam 99 – active site structure 102, 112 – atomic structures 127 – general features 102 – mutagenesis studies 105 PAMO, BVMO 82 Panus tigrinus, laccase 57 paper mill effluent treatment, laccase 56 pathway engineering, steroid biotransformation 170

297

298

Index PDOR, see phthalate dioxygenase reductase peracid catalyst 79 periplasmic oxidase systems 3 peroxide shunt 100, 126, 162, 284 peroxides 142 peroxyflavin enzyme intermediate 81 Pfam database 58 pharma research 239 pharmaceutical biooxidations 233 phase I drug-metabolizing enzymes 212–213 phenazine methosulfate, see PMS phenols 43 2-phenoxypropionic acid (POPS) 199 phenylacetone monooxygenase 83 3-phenyl catechol, bioconversion 199 – 199 2-phenylphenol, bioconversion 199 phosphite dehydrogenase, oxidative biocatalysts 87 photoreaction system, NAD(P)H 271 phthalate dioxygenase reductase (PDOR) 136 phylogenetic tree, type I BVMOs 85 plant protection, laccase 51 plasmids, biotransformation 180–181 plastics degradation, laccase 63 Pleurotus ostreatus, laccase 57 PMS, enzyme activity 5 pollutants, laccase 63 polychlorinated aromatics, CYP 107 polyol oxidation 14 – L-ascorbate 23 – Bertrand–Hudson’s rule 15 – dehydrogenases 17, 23 – dihydroxyacetone 19 – L-erythrulose 16 – GLDH 15 – Gluconobacter 17 – Gluconobacter frateurii 21 – Gluconobacter industrius 15 – Gluconobacter oxydans 23 – Gluconobacter suboxydans 14 – keto-gluconicigenium vulgare 23 – ribitol dehydrogenase 21 – L-ribulose 21 – D -sorbitol dehydrogenase 15, 22 – L-sorbose 22 – L-sorbosone dehydrogenase 23 POPS, see 2-phenoxypropionic acid PQQ (pyrroloquinoline quinone) – dehydrogenases 6 Pravastatin 179, 182 – biotransformation 179, 182

primary dehydrogenases 2 progesterone 160 protein–polyion fi lms 276 pseudocumene 200 Pseudomonas fluorescens, P450-acyl CoA dehydrogenase fusion 143 Pseudomonas putida 99, 124, 169, 194, 201 – alcohol dehydrogenases 9 – CYP 99 Pseudomonas sp. – alcohol dehydrogenase 10 – glucose oxidation 11 putidaredoxin (Pd) 128 putidaredoxin reductase (PdR) 128 pyrroloquinoline quinone, see PQQ

q quinate dehydrogenase (QDH) 24–25 quinohemoprotein 6–7 quinoprotein D -sorbitol dehydrogenase (SLDH) 15 quinoprotein GDHs 13

r radicals 52, 65 rat CPR 129 rat liver P450 167 reactive complex geometry 254 recombinant human CYP (rhCYP) systems 215 recombinant human DMEs 212 recycling, see cofactor regeneration redox partners 123 reductase – adrenodoxin 127–128, 163 – BMVO 78 – CYP 129 – domains 140 – putidaredoxin (Pd) 101, 124 – putidaredoxin reductase (PdR) 101, 129 regeneration, enzymatic 267–271 reoxidization of flavoenzymes 66 Rhizopus arrhizus 160 Rhizopus nigricans 160 Rhodococcus erythropolis 136 Rhodococcus rhodochrous 138 ribitol dehydrogenase 21 L-ribulose 21

s Saccharomyces cerevisiae – drug discovery 238 – drug metabolism 218

Index secondary alcohol dehydrogenase 9 self-sufficient synthesis 170 sensors – alcohol 8 – glucose 13 – see also biosensors sequence clustering, oxidative biocatalysts 84 shikimate pathway, quinate oxidation 24 SNDH1, see L-sorbosone dehydrogenase 1 D -sorbitol dehydrogenase (SLDH) 32 L-sorbose 32 L-sorbosone dehydrogenase 1 (SNDH1) 22 Soret band 124 stainless steel stirred tank bioreactor 248 steroid biotransformations 155–176 – Arabidopsis thaliana 170 – Bacillus megaterium 164 – genetically modified microorganisms 163–170 – pathway engineering 170 steroid hormones 170 steroid hydroxylation 155–176 – CYP dependent 156–159 Streptomyces carbophilus 179 Streptomyces griseus 267 Streptomyces roseochromogenes 162 Streptomyces/Actinomyces 228 styrene, heterocyclic compounds 201 styrene oxide production 200 substituting NAD(P)H, see cofactor regeneration substitution cofactors – electrochemical systems 272–278 – photochemical approaches 271–272 –, see cofactors, substitution Sulfolobus solfataricus 143 superoxide, abortive catalytic cycles 142 surfactant fi lms, electrochemistry 276–278

t taint prevention Tamiflu 25

56

TEMPO 60, 65 terpenoid compounds, CYP 111 testosterone 188 Thermobifi da fusca 83 thermotolerant acetic acid bacteria Trametes villosa 44 Tricholoma giganteum 58 two-liquid phase system 200 two-step biotransformation 203 type I BVMOs – catalytic mechanism 81 – phylogenetic tree 85 – structure 80 – see also Baeyer–Villiger monooxygenases

21–22

u ubiquinones (UQ) 3 uncoupling – P450 biocatalysis – pathways 100

88

w warfare agents 62 Warfarin 258 waste treatment 55 white laccase 57 whole cells 194 – biotransformation 201, 246–247 – catalysts 86, 181–182 wine bottling, laccase 55

x D -xylulose

14

y yeast

233–252 – biotransformations 156 – heterologous expression 236–237 – industrial oxidation 194 – vector 236 yellow laccase 57

299

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