Practical Methods for Biocatalysis and Biotransformations

Practical Methods for Biocatalysis and Biotransformations Editors JOHN WHITTALL Manchester Interdisciplinary Biocentre...

Author: John Whittall | Peter Sutton

189 downloads 2064 Views 5MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!
Report copyright / DMCA form

DOWNLOAD PDF

Practical Methods for Biocatalysis and Biotransformations Editors

JOHN WHITTALL Manchester Interdisciplinary Biocentre, University of Manchester, United Kingdom PETER SUTTON Synthetic Chemistry, GlaxoSmithKline R&D Ltd, United Kingdom

A John Wiley and Sons, Ltd., Publication

Practical Methods for Biocatalysis and Biotransformations

Practical Methods for Biocatalysis and Biotransformations Editors

JOHN WHITTALL Manchester Interdisciplinary Biocentre, University of Manchester, United Kingdom PETER SUTTON Synthetic Chemistry, GlaxoSmithKline R&D Ltd, United Kingdom

A John Wiley and Sons, Ltd., Publication

This edition first published 2010 Ó 2010 John Wiley & Sons Ltd Registered office John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, United Kingdom For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com. The right of the author to be identified as the author of this work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. The publisher and the author make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of fitness for a particular purpose. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for every situation. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of experimental reagents, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each chemical, piece of equipment, reagent, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. The fact that an organization or Website is referred to in this work as a citation and/or a potential source of further information does not mean that the author or the publisher endorses the information the organization or Website may provide or recommendations it may make. Further, readers should be aware that Internet Websites listed in this work may have changed or disappeared between when this work was written and when it is read. No warranty may be created or extended by any promotional statements for this work. Neither the publisher nor the author shall be liable for any damages arising herefrom. Library of Congress Cataloging-in-Publication Data Practical methods for biocatalysis and biotransformations / editors, John Whittall, Peter Sutton. p. ; cm. Includes bibliographical references and index. ISBN 978-0-470-51927-1 1. Enzymes—Biotechnology. 2. Biotransformation (Metabolism) 3. Organic compounds— Synthesis. I. Whittall, John. II. Sutton, Peter (Peter W.) [DNLM: 1. Biocatalysis. 2. Biotransformation. 3. Enzymes. QU 135 P895 2009] TP248.65.E59P73 2009 660.60 34—dc22 2009030811 A catalogue record for this book is available from the British Library. ISBN 978-0-470-51927-1 Set in 10/12pt Times by Integra Software Services Pvt. Ltd, Pondicherry, India Printed and bound in Great Britain by CPI Antony Rowe, Chippenham, Wiltshire

Contents

Preface Abbreviations List of Contributors 1

Biotransformations in Small-molecule Pharmaceutical Development Joseph P. Adams, Andrew J. Collis, Richard K. Henderson and Peter W. Sutton

2

Biocatalyst Identification and Scale-up: Molecular Biology for Chemists Kathleen H. McClean

3

Kinetic Resolutions Using Biotransformations 3.1 Stereo- and Enantio-selective Hydrolysis of rac-2-Octylsulfate Using Whole Resting Cells of Pseudomonas spp. Petra Gadler and Kurt Faber Protease-catalyzed Resolutions Using the 3-(3-Pyridine)propionyl Anchor Group: p-Toluenesulfonamide Christopher K. Savile and Romas J. Kazlauskas 3.3 Desymmetrization of Prochiral Ketones Using Enzymes Andrew J. Carnell

xi xiii xix 1

83

117 117

3.2

3.4

4

Enzymatic Resolution of 1-Methyl-tetrahydroisoquinoline using Candida rugosa Lipase Gary Breen

Dynamic Kinetic Resolution for the Synthesis of Esters, Amides and Acids Using Lipases 4.1 Dynamic Kinetic Resolution of 1-Phenylethanol by Immobilized Lipase Coupled with In Situ Racemization over Zeolite Beta Kam Loon Fow, Yongzhong Zhu, Gaik Khuan Chuah and Stephan Jaenicke

121 125

129

133 133

vi

Contents

4.2

4.3

Synthesis of the (R)-Butyrate Esters of Secondary Alcohols by Dynamic Kinetic Resolution Employing a Bis(tetrafluorosuccinato)bridged Ru(II) Complex S.F.G.M. van Nispen, J. van Buijtenen, J.A.J.M. Vekemans, J. Meuldijk and L.A. Hulshof

137

Dynamic Kinetic Resolution 6,7-Dimethoxy-1-methyl-1,2,3,4tetrahydroisoquinoline Michael Page, John Blacker and Matthew Stirling

141

4.4

Dynamic Kinetic Resolution of Primary Amines with a Recyclable Palladium Nanocatalyst (Pd/AlO(OH)) for Racemization Soo-Byung Ko, Mahn-Joo Kim and Jaiwook Park 4.5 Dynamic Kinetic Resolution of Amines Involving Biocatalysis and In Situ Free-radical-mediated Racemization Ste´phane Gastaldi, Ge´rard Gil and Miche`le P. Bertrand

5

4.6

Chemoenzymatic Dynamic Kinetic Resolution of (S)-Ibuprofen A.H. Kamaruddin and F. Hamzah

4.7

Dynamic Kinetic Resolution Synthesis of a Fluorinated Amino Acid Ester Amide by a Continuous Process Lipase-mediated Ethanolysis of an Azalactone Matthew Truppo, David Pollard, Jeffrey Moore and Paul Devine

Enzymatic Selectivity in Synthetic Methods 5.1 Alcalase-catalysed Syntheses of Hydrophilic Di- and Tri-peptides in Organic Solvents Xue-Zhong Zhang, Rui-Zhen Hou, Li Xu and Yi-Bing Huang 5.2 Selective Alkoxycarbonylation of 1,25-Dihydroxyvitamin D3 Diol Precursor with Candida antarctica Lipase B Miguel Ferrero, Susana Fernańdez and Vicente Gotor The Use of Lipase Enzymes for the Synthesis of Polymers and Polymer Intermediates Alan Taylor 5.4 Bioconversion of 3-Cyanopyridine into Nicotinic Acid with Gordona terrae NDB1165 Tek Chand Bhalla

148

153 157

162

165 165

170

5.3

5.5

Enzyme-promoted Desymmetrization of Prochiral Dinitriles Marloes A. Wijdeven, Piotr Kiełbasin´ski and Floris P.J.T. Rutjes 5.6 Epoxide Hydrolase-catalyzed Synthesis of (R)-3-Benzyloxy-2methylpropane-1,2-diol Takeshi Sugai, Aya Fujino, Hitomi Yamaguchi and Masaya Ikunaka 5.7 One-pot Biocatalytic Synthesis of Methyl (S)-4-Chloro-3hydroxybutanoate and Methyl (S)-4-Cyano-3-hydroxybutanoate Maja Majeric´ Elenkov, Lixia Tang, Bernhard Hauer and Dick B. Janssen

173

182 186

190

199

Contents

6

Aldolase Enzymes for Complex Synthesis 6.1 One-step Synthesis of L-Fructose Using Rhamnulose-1-phosphate Aldolase in Borate Buffer William A. Greenberg and Chi-Huey Wong 6.2

Straightforward Fructose-1,6-bisphosphate Aldolase-mediated Synthesis of Aminocyclitols Marielle Lemaire and Lahssen El Blidi

Synthesis of D-Fagomine by Aldol Addition of Dihydroxyacetone to N-Cbz-3-Aminopropanal Catalysed by D-Fructose-6-phosphateAldolase Jose´ A. Castillo, Teodor Parella, Tomoyuki Inoue, Georg A. Sprenger, Jesu´s Joglar and Pere Clape´s 6.4 Chemoenzymatic Synthesis of 5-Thio-D-xylopyranose Franck Charmantray, Philippe Dellis, Virgil He´laine, Soth Samreth and Laurence Hecquet

vii

203 203

206

6.3

7

Enzymatic Synthesis of Glycosides and Glucuronides 7.1 Glycosynthase-assisted Oligosaccharide Synthesis Adrian Scaffidi and Robert V. Stick 7.2 Glycosyl Azides: Novel Substrates for Enzymatic Transglycosylations Vladimı´r Krˇen and Pavla Bojarova´ 7.3

Facile Synthesis of Alkyl -D-Glucopyranosides from D-Glucose and the Corresponding Alcohols Using Fruit Seed Meals Wen-Ya Lu, Guo-Qiang Lin, Hui-Lei Yu, Ai-Ming Tong and Jian-He Xu

7.4

Laccase-mediated Oxidation of Natural Glycosides Cosimo Chirivı`, Francesca Sagui and Sergio Riva 7.5 Biocatalysed Synthesis of Monoglucuronides of Hydroxytyrosol, Tyrosol, Homovanillic Alcohol and 3-(40 -Hydroxyphenyl)propanol Using Liver Cell Microsomal Fractions Olha Khymenets, Pere Clape´s, Teodor Parella, Marıá-Isabel Covas, Rafael de la Torre, and Jesu´s Joglar 7.6 Synthesis of the Acyl Glucuronide of Mycophenolic Acid Matthias Kittelmann, Lukas Oberer, Reiner Aichholz and Oreste Ghisalba 8

Synthesis of Cyanohydrins Using Hydroxynitrile Lyases 8.1 Synthesis of (S)-2-Hydroxy-2-methylbutyric Acid by a Chemoenzymatic Methodology Manuela Avi and Herfried Griengl 8.2 (S)-Selective Cyanohydrin Formation from Aromatic Ketones Using Hydroxynitrile Lyases Chris Roberge, Fred Fleitz and Paul Devine

212

218

227 227

232

236

240

245

251

255 255

259

viii

Contents

8.3

Hydroxynitrile-lyase-catalysed Synthesis of Enantiopure (S)-Acetophenone Cyanohydrins Jan von Langermann, Annett Mell, Eckhard Paetzold and Udo Kragl 8.4 (R)- and (S)-Cyanohydrin Formation from Pyridine3-carboxaldehyde Using CLEATM-immobilized Hydroxynitrile Lyases Chris Roberge, Fred Fleitz and Paul Devine

8.5

9

A New (R)-Hydroxynitrile Lyase from Prunus mume for Asymmetric Synthesis of Cyanohydrins Yasuhisa Asano

Synthesis of Chiral sec-Alcohols by Ketone Reduction 9.1 Asymmetric Synthesis of (S)-Bis(trifluoromethyl)phenylethanol by Biocatalytic Reduction of Bis(trifluoromethyl)acetophenone David Pollard, Matthew Truppo and Jeffrey Moore 9.2 Enantioselective and Diastereoselective Enzyme-catalyzed Dynamic Kinetic Resolution of an Unsaturated Ketone Birgit Kosjek, David Tellers and Jeffrey Moore 9.3

Enzyme-catalysed Synthesis of -Alkyl- -hydroxy Ketones and Esters by Isolated Ketoreductases Ioulia Smonou and Dimitris Kalaitzakis

Asymmetric Reduction of Phenyl Ring-containing Ketones Using Xerogel-encapsulated W110A Secondary Alcohol Dehydrogenase from Thermoanaerobacter ethanolicus Musa M. Musa, Karla I. Ziegelmann-Fjeld, Claire Vieille, J. Gregory Zeikus and Robert S. Phillips 9.5 (R)- and (S)-Enantioselective Diaryl Methanol Synthesis Using Enzymatic Reduction of Diaryl Ketones Matthew Truppo, Krista Morley, David Pollard and Paul Devine

262

266

269

273 273

276

278

9.4

9.6

10

Highly Enantioselective and Efficient Synthesis of Methyl (R)-o-Chloromandelate, Key Intermediate for Clopidogrel Synthesis, with Recombinant Escherichia coli Tadashi Ema, Nobuyasu Okita, Sayaka Ide and Takashi Sakai

284

288

291

Reduction of Functional Groups 10.1 Reduction of Carboxylic Acids by Carboxylic Acid Reductase Heterologously Expressed in Escherichia coli Andrew S. Lamm, Arshdeep Khare and John P.N. Rosazza

295

10.2

Light-driven Stereoselective Biocatalytic Oxidations and Reductions Andreas Taglieber, Frank Schulz, Frank Hollmann, Monika Rusek and Manfred T. Reetz

299

10.3

Unnatural Amino Acids by Enzymatic Transamination: Synthesis of Glutamic Acid Analogues with Aspartate Aminotransferase Thierry Gefflaut, Emmanuelle Sagot and Jean Bolte

295

306

Contents

Synthesis of L-Pipecolic Acid with 1-Piperidine-2-carboxylate Reductase from Pseudomonas putida Hisaaki Mihara and Nobuyoshi Esaki 10.5 Synthesis of Substituted Derivatives of L-Phenylalanine and of other Non-natural L-Amino Acids Using Engineered Mutants of Phenylalanine Dehydrogenase Philip Conway, Francesca Paradisi and Paul Engel

ix

10.4

11

310

314

Enzymatic Oxidation Chemistry 11.1 Monoamine Oxidase-catalysed Reactions: Application Towards the Chemo-enzymatic Deracemization of the Alkaloid (–)-Crispine A Andrew J. Ellis, Renate Reiss, Timothy J. Snape and Nicholas J. Turner

319

11.2

323

Glucose Oxidase-catalysed Synthesis of Aldonic Acids Fabio Pezzotti, Helene Therisod and Michel Therisod 11.3 Oxidation and Halo-hydroxylation of Monoterpenes with Chloroperoxidase from Leptoxyphium fumago Bjoern-Arne Kaup, Umberto Piantini, Matthias Wu¨st and Jens Schrader 11.4 Chloroperoxidase-catalyzed Oxidation of Phenyl Methylsulfide in Ionic Liquids Cinzia Chiappe 11.5

Stereoselective Synthesis of -Hydroxy Sulfoxides Catalyzed by Cyclohexanone Monooxygenase Stefano Colonna, Nicoletta Gaggero, Sara Pellegrino and Francesca Zambianchi

Enantioselective Kinetic Resolution of Racemic 3-Phenylbutan-2one Using a Baeyer–Villiger Monooxygenase Anett Kirschner and Uwe T. Bornscheuer 11.7 Desymmetrization of 1-Methylbicyclo[3.3.0]octane-2,8-dione by the Retro-claisenase 6-Oxo Camphor Hydrolase Gideon Grogan and Cheryl Hill

319

327

330

332

11.6

11.8

12

Synthesis of Optically Pure Chiral Lactones by Cyclopentadecanone Monooxygenase-catalyzed Baeyer–Villiger Oxidations Shaozhao Wang, Jianzhong Yang and Peter C.K. Lau

Whole-cell Oxidations and Dehalogenations 12.1 Biotransformations of Naphthalene to 4-Hydroxy-1-tetralone by Streptomyces griseus NRRL 8090 Arshdeep Khare, Andrew S. Lamm and John P.N. Rosazza 12.2 Hydroxylation of Imidacloprid for the Synthesis of Olefin Imidacloprid by Stenotrophomonas maltophilia CGMCC 1.1788 Sheng Yuan and Yi-jun Dai

337

341

344

351 351

355

x

Contents

12.3

Biocatalytic Synthesis of 6-Hydroxy Fluvastatin using Motierella rammaniana DSM 62752 in Shake Flask Culture and on Multi-gram Scale using a Wave Bioreactor Matthias Kittelmann, Maria Serrano Correia, Anton Kuhn, Serge Parel, Ju¨rgen Ku¨hno¨l, Reiner Aichholz, Monique Ponelle and Oreste Ghisalba 12.4 Synthesis of 1-Adamantanol from Adamantane through Regioselective Hydroxylation by Streptomyces griseoplanus Cells Koichi Mitsukura, Yoshinori Kondo, Toyokazu Yoshida and Toru Nagasawa 12.5

Enantioselective Benzylic Microbial Hydroxylation of Indan and Tetralin Renata P. Limberger, Cleber V. Ursini, Paulo J.S. Moran and J. Augusto R. Rodrigues

Stereospecific Biotransformation of (R,S)-Linalool by Corynespora cassiicola DSM 62475 into Linalool Oxides Marco-Antonio Mirata and Jens Schrader 12.7 The Biocatalytic Synthesis of 4-Fluorocatechol from Fluorobenzene Louise C. Nolan and Kevin E. O’Connor

359

367

369

12.6

Synthesis of Enantiopure (S)-Styrene Oxide by Selective Oxidation of Styrene by Recombinant Escherichia coli JM101 (pSPZ10) Katja Buehler and Andreas Schmid 12.9 Biotransformation of -Bromo and ,0 -Dibromo Alkanone into -Hydroxyketone and -Diketone by Spirulina platensis Takamitsu Utsukihara and C. Akira Horiuchi

376 379

12.8

Index

385

391

397

Preface

During the early to mid 1990s Professor Stan Roberts was chief editor of a series of looseleaf laboratory protocols detailing the use of biotransformations in synthetic organic chemistry that were collected together and published in book form (Preparative Biotransformations, Wiley, Chichester, 1999). This led to the publication of the series of books Catalysts for Fine Chemical Synthesis, volumes 1–5, by the same publisher which covered the application of chemo- and bio-catalytic procedures for the synthesis of fine chemicals; for this series, Dr John Whittall became co-editor on the homogeneous catalysis volumes. Following the format of this series, Practical Methods in Biocatalysis and Biotransformations has been prepared. In keeping with these earlier formats, we aim to provide the readership with enough information to understand when a biocatalytic or biotransformation method would be a suitable practical method to carry out their synthetic transformation. In recent times, the employment of enzymes and whole cells to perform a range of organic reactions has become much more commonplace, and biotransformation has become accepted as a powerful method for application in synthetic organic chemistry. However, for chemists developing synthetic methods for a particular target molecule, the understanding of the advantages and limitations of biocatalysis and biotransformation is not always clear. Therefore, this book intends to review the industrial background to when biotransformations are used and introduce the nonmicrobiologist to the background of how biocatalysts are discovered and developed and then give detailed experimental procedures for a comprehensive range of useful biotransformation methods. In order to place the later chapters in proper context, Chapter 1 offers a comprehensive review of biotransformation from the perspective of a large pharmaceutical company (GSK) and Chapter 2 gives an introduction that allows an appreciation of molecular biology for scientists with no formal training in this area. In the remaining chapters, key biotransformations have been identified from the recent primary literature (learned journals) and the respective authors have amplified the disclosure of their methodologies in this volume. These disclosures often contain additional equipment and experimental details to those found in the experimental section of most journals, allowing the reader to decide whether these methods are suitable for addressing their needs. Chapter 3 describes the application of lipases, proteases and sulfatases for the kinetic resolution of a range of interesting molecules. A selection of dynamic kinetic resolution (DKR) procedures is disclosed in Chapter 4. DKRs are attracting a significant amount of

xii

Preface

interest as they allow access to >50 % yields of single enantiopure products from racemates. Other useful synthetic applications of hydrolase enzymes are covered in Chapter 5, including desymmetrization and regio- and chemo-selective transformations. Chapters 6 and 7 cover sugar-type chemistry, focusing on aldol and glycosylation methods which can offer substantial advantages over traditional chemical approaches. Chapter 8 describes the application of hydroxyl nitrile lyases to the synthesis of new chiral cyanohydrins and -hydroxy acids and includes new approaches to the transformation of ‘difficult’ aldehyde and ketone substrates using substrate engineering and immobilization techniques. The latter part of the book is dedicated to redox biotransformation application, with Chapter 9 disclosing several methods for the synthesis of chiral secondary alcohols using a range of commercially available ketoreductases (alcohol dehydrogenases) which are now being applied regularly on a large scale. Chapter 10 covers reductive enzymes with an emphasis on transaminase enzymes, which are enjoying widespread application in the synthesis of nonnatural amino acids which are key building blocks for several products of industrial importance. The use of a range of oxidative enzymes in synthesis is covered in Chapter 11, whilst the very powerful technique of regio- and stereo-specific biohydroxylation of even complex molecules by fermenting whole-cell methods is covered in Chapter 12. The Editors are most grateful to the authors who have submitted details of their procedures in the prescribed format for inclusion in this book. We hope that this book will increase the exposure of these methods to the chemical community and contribute to the expanded employment of biocatalysis in organic synthesis. John Whittall, Manchester Peter Sutton, Stevenage 2009

Abbreviations

A ABTS 7-ACA ACN AcOH ACS GCI 7-ADCA ADH ADH-RE AIBN 6-APA API Ara-G Ara-U AspAT AT AZT BEHP BES BLAST BREP BSA BSA BVMO C CAL-A CAL-B Car CASTing Cbz CCL

adenine 2,20 -azino-bis-3-ethylbenzothiazoline-6-sulfonic acid 7-aminocephalosporanic acid acetonitrile acetic acid American Chemical Society Green Chemistry Institute 7-aminodesacetoxycephalosporanic acid alcohol dehydrogenase (alternative name for a ketoreductases or KREDs) alcohol dehydrogenase from Rhodococcus erythropolis 2,20 -azobis(2-methylpropionitrile) 6-aminopenicillanic acid active pharmaceutical ingredient 9-b-D-arabinofuranosylguanidine 9-b-D-arabinofuranosyluridine aspartate aminotransferase aminotransferases 30 -azido-20 ,30 -dideoxythymidine (zidovudine) bis(2-ethylhexyl)phthalate N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid basic local alignment search tool butanol-rinsed enzyme preparation bovine serum albumin N-bromosuccinimide Baeyer–Villiger monooxygenase cytosine lipase A from Candida antarctica lipase B from Candida antarctica carboxylic acid reductase combinatorial active site saturation test Benzyloxycarbonyl lipase from Candida cylindracea (now known as lipase from Candida rugosa or CRL)

xiv

Abbreviations

CDI CDW cGMP CHMO CINV CLEA CLEC CNS CPDMO CPO CRL CSA CYP DBDMH DBE DCM DCW DDI DERA dGTP DHA DHAP DHF DIPE DKR DMAP DMF DMSO DNA DNAse dNTP DoE DOT DSMZ dsDNA D4T DTT dUDP dUMP E EDC EDTA

1,10 -carbonyldiimidazole cell dry weight current good manufacturing practice cyclohexanone monooxygenase chemotherapy-induced nausea cross-linked enzyme aggregate cross-linked enzyme crystal Central nervous system cyclopentadecanone monooxygenase chloroperoxidase lipase from Candida rugosa cysteine sulfinic acid cytochrome P450 N,N0 -dibromodimethylhydantoin di-n-butylether dichloromethane dry cell weight drug–drug interaction 2-deoxyribose-5-phosphate aldolase deoxyguanosine triphosphate dihydroxyacetone dihydroxyacetone phosphate dihydrofolate diisopropylether dynamic kinetic resolution 4-dimethylaminopyridine dimethylformamide dimethylsulfoxide deoxyribonucleic acid deoxyribonuclease deoxyribonucleotide triphosphate design of experiment dissolved oxygen tension Deutsche Sammlung von Mikroorganismen und Zellkulturen double-stranded DNA dideoxydidehydrothymidine dithiothreitol 20 -deoxyuridine-50 -diphosphate 20 -deoxyuridine-50 -monophosphate enantiomeric ratio 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride ethylenediaminetetraacetic acid

Abbreviations

EHS epPCR EtOAc FACS FAD FADH2 FASTA FDA FDH FMN FPLC FruA FSA FTIR GABA G GC GDH GlcI GMO GMM G6P G6PDH GPC GPO GR GRAS GSK HBV HEPES HIV HMQC HNL HOBt HOPhPr HOTYR HPA HPI HPLC HTS HVAlc

environmental health and safety error-prone PCR Ethyl acetate fluorescence-activated cell sorting flavin adenine dinucleotide flavin adenine dinucleotide, reduced form FAST ALL (a programme for fast protein comparison or fast nucleotide sequence comparison) Food and Drug Administration (United States) formate dehydrogenase flavin mononucleotide (riboflavin-50 -phosphate) fast protein liquid chromatography fructose-1,6-bisphosphate aldolase D-fructose-6-phosphate aldolase Fourier-transform infrared spectroscopy g-aminobutyric acid guanine gas chromatography glucose dehydrogenase glucose isomerase genetically modified organism genetically modified microorganism glucose-6-phosphate glucose-6-phosphate dehydrogenase gel permeation chromatography L-glycerol-3-phosphate oxidase glucocorticoid receptor generally recognized as safe GlaxoSmithKline hepatitis B virus 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid human immunodeficiency virus heteronuclear multiple quantum coherence hydroxynitrile lyase 1-hydroxybenzotriazole hydroxyphenylpropanol hydroxytyrosol hydroxypyruvate N-hydroxyphthalimide high-performance liquid chromatography high-throughput screening Homovanillic alcohol

xv

xvi

Abbreviations

IMI Indels IP IPTG ISPR KPB KR KRED LAS LB LCA LovD Mab MAO-N MEA MES MGF MML MOPS m.p. MPA MPLC mRNA MS MTBE MTQ MYB NADþ NADH NADPH NADPþ NAG NAM NANA NCE NK-1 NME NMR NP OCH ORI P450

imidacloprid insertions and deletions intellectual property isopropyl-b-D-thiogalactopyranoside in situ product removal potassium phosphate buffer kinetic resolution ketoreductase (alternative name for an alcohol dehydrogenase or ADH) lovastatin ammonium salt Luria–Bertani life cycle analysis acyltransferase from the lovastatin biosynthetic pathway monoclonal antibody monoamine oxidase malt extract agar 2-morpholino ethansulfonic acid monohydrate minimum genome factories lipase from Mucor sp. 3-morpholino propane sulfonic acid melting point mycophenolic acid medium-pressure chromatography messenger RNA molecular sieves tert-butylmethylether methyl-tetrahydroisoquinoline malt yeast broth b-nicotinamide adenine dinucleotide b-nicotinamide adenine dinucleotide, reduced form b-nicotinamide adenine dinucleotide 20 -phosphate, reduced form b-nicotinamide adenine dinucleotide 20 -phosphate N-acetyl-D-glucosamine N-acetyl-D-mannosamine N-acetyl-D-neuraminic acid new chemical entity neurokinin-1 new molecular entity nuclear magnetic resonance nucleoside phosphorylase 6-oxo camphor hydrolase origin of replication cytochrome P450

Abbreviations

P450 BM-3 PAMO Pase PAT PCL PCR PDCB PEP PFL PGA Pip2C PLE pNPG PNP PPL ProSAR QbD QSAR RAMA R&D rDNA rRNA Rf RhaD RNA ROH Rt SAS SCR SIGEX SMB SOT ssDNA T Taq TBDMSCl TBME TCA TDP TdR TEMPO

cytochrome P450 BM-3 from Bacillus megaterium phenylacetone monooxygenase acid phosphatase process analytical technology lipase from Pseudomonas cepacia (now renamed to Burkholderia cepacia) polymerase chain reaction potato–dextrose–carrot broth phosphoenolpyruvic acid lipase from Pseudomonas fluorescens penicillin G acylase D1-piperideine-2-carboxylate reductase pig liver esterase p-nitrophenyl-b-D-glucopyranoside purine nucleoside phosphorylase porcine pancreatic lipase protein sequence–activity relationship quality by design quantitative structure–activity relationship rabbit muscle aldolase (fructose-1,6-bis-phosphate aldolase) research and development recombinant DNA ribosomal RNA retention factor rhamnulose-1-phosphate aldolase ribonucleic acid generic alcohol retention time simvastatin ammonium salt Saccharomyces cerevisiae carbonyl reductase substrate-induced gene-expression screening simulated moving bed chromatography Spirulina–Ogawa–Terui single-stranded DNA thymine a thermostable DNA polymerase from Thermus aquaticus tert-butyldimethylsilyl chloride tert-butylmethylether trichloroacetic acid thymidine 50 -phosphate thymidine 2,2,6,6-tetramethyl-1-piperidinyloxy

xvii

xviii

Abbreviations

TFA THF THFo ThDP TK TLC TMP TMS TMOS TMSOTf TP tris-HCl TTN TYR U U UdR UDP UTP UDPGA UDPGT URDP UV VVM WFCC YPG

trifluoroacetic acid tetrahydrofuran tetrahydrofolate thiamine pyrophosphate transketolase thin-layer chromatography thymidine 50 -monophosphate tetramethyl silane tetramethyl orthosilicate trimethylsilyl triflate thymidine-50 -phosphorylase tris(hydroxymethyl)aminomethane HCl total turnover number tyrosol unit of enzyme activity (mmol min1) uracil 20 -deoxyuridine uridine-50 -diphosphate uridine-50 -triphosphate uridine-50 -diphosphoglucuronic acid uridine-50 -diphosphoglucuronyl transferase uridine-50 -phosphorylase ultraviolet gas volume flow per unit of liquid volume per minute World Federation for Culture Collections yeast extract–peptone–glucose

List of Contributors

Joseph P. Adams, GlaxoSmithKline, Synthetic Chemistry, Gunnels Wood Road, Stevenage, Hertfordshire SG1 2NY, UK Reiner Aichholz, Metabolism and Pharmacokinetics, NIBR, Novartis Pharma AG, CH-4002 Basel, Switzerland Yasuhisa Asano, Biotechnology Research Center and Department of Biotechnology, Toyama Prefectural University, 5180 Kurokawa, Imizu, Toyama 939-0398, Japan Manuela Avi, Institute of Organic Chemistry, Graz University of Technology, Stremayrgasse 16, 8010 Graz, Austria Miche`le P. Bertrand, Laboratoire de Chimie Molećulaire Organique, LCP UMR 6264, Boite 562, Universite´ Paul Ce´zanne, Aix-Marseille III, Faculte´ des Sciences St Je´roˆme, Avenue Escadrille Normandie-Niemen, 13397 Marseille Cedex 20, France Tek Chand Bhalla, Department of Biotechnology, Himachal Pradesh University, Shimla 171005, India John Blacker, NPIL Pharma Ltd, Leeds Road, Huddersfield, HD1 9GA, UK Lahssen El Blidi, Laboratoire SEESIB, UMR 6504 CNRS, Universite´ Blaise Pascal, 24 avenue des Landais 63177 Aubie`re cedex, France Pavla Bojarova´, Institute of Microbiology, Center of Biocatalysis and Biotransformations, Academy of Sciences of the Czech Republic, Vı´denˇska´ 1083, CZ-142 20 Prague 4, Czech Republic Jean Bolte, Department of Chemistry, Universite´ Blaise Pascal, Clermont-Ferrand, France Uwe T. Bornscheuer, Department of Biotechnology and Enzyme Catalysis, Institute of Biochemistry, Greifswald University, Felix-Hausdorff-Str. 4, 17487 Greifswald, Germany Gary Breen, GlaxoSmithKline, Synthetic Chemistry, Leigh, Tonbridge, Kent, TN11 9AN, UK Katja Buehler, Laboratory of Chemical Biotechnology, Faculty of Biochemical and Chemical Engineering, TU Dortmund, Emil-Figge-Strasse 66, 44221 Dortmund, Germany

xx


J. van Buijtenen, Eindhoven University of Technology, Laboratory of Macromolecular and Organic Chemistry, PO Box 513, 5600 MB Eindhoven, The Netherlands Andrew J. Carnell, Department of Chemistry, Robert Robinson Laboratories, University of Liverpool, Liverpool, L69 7ZD, UK Jose´ A. Castillo, Biotransformation and Bioactive Molecules Group, Instituto de Quimica Avanzada de Catalunã, Consejo Superior de Investigaciones Cientificas, Jordi Girona 18-26, 08034 Barcelona, Spain Franck Charmantray, Laboratoire SEESIB, UMR 6504 CNRS, Universite´ Blaise Pascal, 24 avenue des Landais, 63177 Aubie`re, France Cinzia Chiappe, Dipartimento di Chimica e Chimica Industriale, Universit di Pisa, 56126 Pisa, Italy Cosimo Chirivı´, Istituto di Chimica del Riconoscimento Molecolare, C.N.R., Via Mario Bianco 9, 20131 Milano, Italy Gaik Khuan Chuah, Department of Chemistry, National University of Singapore, Kent Ridge, Singapore 119260, Republic of Singapore Pere Clape´s, Biotransformation and Bioactive Molecules Group, Instituto de Quimica Avanzada de Catalunã, Consejo Superior de Investigaciones Cientificas, Jordi Girona 18-26, 08034 Barcelona, Spain Andrew J. Collis, GlaxoSmithKline, Biotechnology and Environmental Shared Service, North Lonsdale Road, Ulverston, Cumbria LA12 9DR, UK Stefano Colonna, Dipartimento di Scienze Molecolari Applicate ai Biosistemi (DISMAB), Facolta` di Farmacia, Universita` degli Studi di Milano, via Venezian 21, 20133 Milano, Italy Philip Conway, School of Biomolecular and Biomedical Science, University College Dublin, Belfield, Dublin 4, Ireland Maria Serrano Correia, Rua Maria Auxiliadora, n°147, 6°andar porta 3, Bairro do Rosa´rio, P-2750-616 Cascais, Portugal Marıá-Isabel Covas, Research Unit on Lipids and Cardiovascular Epidemiology, Institut Municipal d’Investigacio´ Me´dica (IMIM). Universitat Pompeu Fabra (CEXS-UPF), Barcelona, Spain Yi-jun Dai, Nanjing Engineering Research Center for microbiology, Jiangsu Key Laboratory for Biodiversity and Biotechnology, College of Life Science, Nanjing Normal University, 1, Wenyuan Rd, Nanjing 210046, PR China Philippe Dellis, Synkem, 47 rue de Longvic, 21300 Chenoˆve, France Paul Devine, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA


xxi

Andrew J. Ellis, School of Chemistry, Manchester Interdisciplinary Biocentre, University of Manchester, 131 Princess Street, Manchester, M1 7DN, UK Tadashi Ema, Division of Chemistry and Biochemistry, Graduate School of Natural Science and Technology, Okayama University, Tsushima, Okayama 700-8530, Japan Paul Engel, School of Biomolecular and Biomedical Science, University College Dublin, Belfield, Dublin 4, Ireland Nobuyoshi Esaki, Institute for Chemical Research, Kyoto University, Uji, Kyoto 611-0011, Japan Kurt Faber, Department of Chemistry, Organic and Bioorganic Chemistry, University of Graz, Heinrichstrasse 28, 8010 Graz, Austria Susana Fernańdez, Departamento de Quı´mica Orgańica e Inorgańica and Instituto Universitario de Biotecnologıá de Asturias, Universidad de Oviedo, 33006-Oviedo (Asturias), Spain Miguel Ferrero, Departamento de Quı´mica Orgańica e Inorgańica and Instituto Universitario de Biotecnologıá de Asturias, Universidad de Oviedo, 33006-Oviedo (Asturias), Spain Fred Fleitz, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA Kam Loon Fow, Department of Chemistry, National University of Singapore, Kent Ridge, Singapore 119260, Republic of Singapore Aya Fujino, Department of Chemistry, Faculty of Science and Technology, Keio University, Hiyoshi, Kohoku-ku, Yokohama 223-8522, Japan Petra Gadler, Department of Chemistry, Organic and Bioorganic Chemistry, University of Graz, Heinrichstrasse 28, 8010 Graz, Austria Nicoletta Gaggero, Dipartimento di Scienze Molecolari Applicate ai Biosistemi (DISMAB), Facolta` di Farmacia, Universita` degli Studi di Milano, via Venezian 21, 20133 Milano, Italy Ste´phane Gastaldi, Laboratoire de Chimie Molećulaire Organique, LCP UMR 6264, Boite 562, Universite´ Paul Ce´zanne, Aix-Marseille III, Faculte´ des Sciences St Je´roˆme, Avenue Escadrille Normandie-Niemen, 13397 Marseille Cedex 20, France Thierry Gefflaut, Department of Chemistry, Universite´ Blaise Pascal, Clermont-Ferrand, France Oreste Ghisalba, Ghisalba Life Sciences GmbH, Habshagstrasse 8c, CH-4153 Reinach, Switzerland Ge´rard Gil, Laboratoire de Ste´reóchimie Dynamique et Chiralite´, ISM2, UMR 6263, Universite´ Paul Ce´zanne, Aix-Marseille III, Faculte´ des Sciences St Je´roˆme, Avenue Escadrille Normandie-Niemen, 13397 Marseille Cedex 20, France

xxii


Vicente Gotor, Departamento de Quı´mica Orgańica e Inorgańica and Instituto Universitario de Biotecnologıá de Asturias, Universidad de Oviedo, 33006-Oviedo (Asturias), Spain William A. Greenberg, Department of Chemistry, The Scripps Research Institute, 10550 North Torrey Pines Rd., La Jolla, CA 92307, USA Herfried Griengl, Research Centre Applied Biocatalysis, Petersgasse 14, 8010 Graz, Austria Gideon Grogan, York Structural Biology Laboratory, Department of Chemistry, University of York, Heslington, York, YO10 5YW, UK F. Hamzah, School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300, Nibong Tebal, Penang, Malaysia Bernhard Hauer, Institute of Technical Biochemistry, University of Stuttgart, Allmandring 31, 70569 Stuttgart, Germany Laurence Hecquet, Laboratoire SEESIB, UMR 6504 CNRS, Universite´ Blaise Pascal, 24 avenue des Landais, 63177 Aubie`re, France Virgil He´laine, Laboratoire SEESIB, UMR 6504 CNRS, Universite´ Blaise Pascal, 24 avenue des Landais, 63177 Aubie`re, France Richard K. Henderson, GlaxoSmithKline, Centre of Excellence for Sustainability and Environment, Park Road, Ware, Hertfordshire SG12 0DP, UK Cheryl Hill, York Structural Biology Laboratory, Department of Chemistry, University of York, Heslington, York, YO10 5YW, UK Frank Hollmann, Max-Planck-Institut fu¨r Kohlenforschung, Kaiser-Wilhelm-Platz 1, 45470 Mu¨lheim/Ruhr, Germany C. Akira Horiuchi, Department of Chemistry, Rikkyo (St.Paul’s) University, NishiIkebukuro, Toshima-Ku, Tokyo 171-8501, Japan Rui-Zhen Hou, Key Laboratory for Molecular Enzymology and Engineering of Ministry of Education, Jilin University, Changchun, 130021, PR China Yi-Bing Huang, Key Laboratory for Molecular Enzymology and Engineering of Ministry of Education, Jilin University, Changchun, 130021, PR China L.A. Hulshof, Eindhoven University of Technology, Laboratory of Macromolecular and Organic Chemistry, PO Box 513, 5600 MB Eindhoven, The Netherlands Sayaka Ide, Division of Chemistry and Biochemistry, Graduate School of Natural Science and Technology, Okayama University, Tsushima, Okayama 700-8530, Japan Masaya Ikunaka, Fine Chemicals Department, Nagase & Co., Ltd., 5-1, NihonbashiKobunacho, Chuo-ku, Tokyo 103-8355, Japan Tomoyuki Inoue, Institute of Microbiology, University of Stuttgart, Allmandring 31, 70569 Stuttgart, Germany Stephan Jaenicke, Department of Chemistry, National University of Singapore, Kent Ridge, Singapore 119260, Republic of Singapore


xxiii

Dick B. Janssen, Biochemical Laboratory, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Nijenborgh 4, 9747 AG, Groningen, The Netherlands Jesu´s Joglar, Biotransformation and Bioactive Molecules Group, Instituto de Quimica Avanzada de Catalunã, Consejo Superior de Investigaciones Cientificas, Jordi Girona 18-26, 08034 Barcelona, Spain Dimitris Kalaitzakis, Department of Chemistry, University of Crete, Iraklion-Voutes, 71003 Crete, Greece Azlina Kamaruddin, School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300, Nibong Tebal, Penang, Malaysia Bjoern-Arne Kaup, DECHEMA e.V., Karl-Winnacker-Institut, Engineering Group, Theodor-Heuss-Allee 25, 60486 Frankfurt, Germany

Biochemical

Romas J. Kazlauskas, Department of Biochemistry, Molecular Biology & Biophysics and The Biotechnology Institute, University of Minnesota, 1479 Gortner Avenue, Saint Paul, MN 55108, USA Arshdeep Khare, Center for Biocatalysis and Bioprocessing, 2501 Crosspark Road, Suite C100 MTF, University of Iowa, Iowa City, Iowa, IA 52242-5000, USA Olha Khymenets, Pharmacology Research Unit, Institut Municipal d’Investigacio´ Me´dica (IMIM), Barcelona, Spain Piotr Kiełbasin´ski, Institute for Molecules and Materials, Radboud University Nijmegen, Toernooiveld 1, NL-6525 ED Nijmegen, The Netherlands Mahn-Joo Kim, Department of Chemistry, Pohang University of Science and Technology (POSTECH), San-31, Hyojadong, Pohang 790-784, Korea Anett Kirschner, Department of Biotechnology and Enzyme Catalysis, Institute of Biochemistry, Greifswald University, Felix-Hausdorff-Str. 4, 17487 Greifswald, Germany Matthias Kittelmann, GDC/PSB/Bioreactions, Novartis Institutes of Biomedical Research (NIBR), Novartis Pharma AG, CH-4002 Basel, Switzerland Soo-Byung Ko, Department of Chemistry, Pohang University of Science and Technology (POSTECH), San-31, Hyojadong, Pohang 790-784, Korea Yoshinori Kondo, Department of Biomolecular Science, Gifu University, Yanagido 1-1, Gifu 501-1193, Japan Birgit Kosjek, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA Udo Kragl, Institut fu¨r Chemie, Universita¨t Rostock, Albert-Einstein-Str. 3a, 18059 Rostock, Germany

xxiv


Vladimı´r Krˇen, Institute of Microbiology, Center of Biocatalysis and Biotransformations, Academy of Sciences of the Czech Republic, Vı´denˇska´ 1083, CZ-142 20 Prague 4, Czech Republic Anton Kuhn, GDC/PSB/Bioreactions, Novartis Institutes of Biomedical Research (NIBR), Novartis Pharma AG, CH-4002 Basel, Switzerland Ju¨rgen Ku¨hno¨l, GDC/PSB/Separations, NIBR, Novartis Pharma AG, CH-4002 Basel, Switzerland Andrew S. Lamm, Center for Biocatalysis and Bioprocessing, 2501 Crosspark Road, Suite C100 MTF, University of Iowa, Iowa City, Iowa, IA 52242-5000, USA Jan von Langermann, Max-Planck-Institut fu¨r Dynamik komplexer technischer Systeme, Physikalisch-Chemische Grundlagen der Prozesstechnik, Sandtorstr.1 D-39106 Magdeburg, Germany Peter C.K. Lau, Biotechnology Research Institute, National Research Council Canada, Montreal, Quebec H4P 2R2, Canada Marielle Lemaire, Laboratoire SEESIB, UMR 6504 CNRS, Universite´ Blaise Pascal, 24 avenue des Landais 63177 Aubie`re cedex, France Renata P. Limberger, State University of Campinas, Institute of Chemistry, CP 6154, 13084-971, Campinas-SP, Brazil Guo-Qiang Lin, Laboratory of Biocatalysis and Bioprocessing, State Key Laboratory of Bioreactor Engineering, East China University of Science and Technology, Shanghai 200237, PR China Wen-Ya Lu, Laboratory of Biocatalysis and Bioprocessing, State Key Laboratory of Bioreactor Engineering, East China University of Science and Technology, Shanghai 200237, PR China Maja Majeric´ Elenkov, Laboratory for Stereoselective Catalysis and Biocatalysis, Ruder Bosˇkovic´ Institute, Bijenicˇka c. 54, 10002 Zagreb, Croatia Kathleen H. McClean, C-Tech Innovation Ltd, Capenhurst Technology Park, Capenhurst, Chester, CH1 6EH, UK Annett Mell, Institut fu¨r Chemie, Universita¨t Rostock, Albert-Einstein-Str. 3a, 18059 Rostock, Germany J. Meuldijk, Eindhoven University of Technology, Laboratory of Macromolecular and Organic Chemistry, PO Box 513, 5600 MB Eindhoven, The Netherlands Hisaaki Mihara, Department of Biotechnology, Institute of Science and Engineering, College of Life Sciences, Ritsumeikan University, Kusatsu, Shiga 525-8577, Japan Marco-Antonio Mirata, DECHEMA e.V., Karl-Winnacker-Institut, Biochemical Engineering Group, Theodor-Heuss-Allee 25, 60486 Frankfurt, Germany


xxv

Koichi Mitsukura, Department of Biomolecular Science, Gifu University, Yanagido 1-1, Gifu 501-1193, Japan Jeffrey Moore, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA Paulo J. S. Moran, State University of Campinas, Institute of Chemistry, CP 6154, 13084-971, Campinas-SP, Brazil Krista Morley, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA Musa M. Musa, Department of Chemistry and of Biochemistry and Molecular Biology, University of Georgia, Athens, GA 30602, USA Toru Nagasawa, Department of Biomolecular Science, Gifu University, Yanagido 1-1, Gifu 501-1193, Japan S.F.G.M. van Nispen, Eindhoven University of Technology, Laboratory of Macromolecular and Organic Chemistry, PO Box 513, 5600 MB Eindhoven, The Netherlands Louise C. Nolan, School of Biomolecular and Biomedical Science, Conway Institute for Biomolecular and Biomedical Research, National University of Ireland, University College Dublin, Ardmore House, Belfield, Dublin 4, Republic of Ireland Kevin E. O’Connor, School of Biomolecular and Biomedical Science, Conway Institute for Biomolecular and Biomedical Research, National University of Ireland, University College Dublin, Ardmore House, Belfield, Dublin 4, Republic of Ireland Lukas Oberer, Analytical and Imaging Sciences, Novartis Institutes of Biomedical Research, Novartis Pharma AG, CH-4002 Basel, Switzerland Nobuyasu Okita, Division of Chemistry and Biochemistry, Graduate School of Natural Science and Technology, Okayama University, Tsushima, Okayama 700-8530, Japan Eckhard Paetzold, Leibniz-Institut fu¨r Katalyse, A.-Einstein-Str. 29a,18059 Rostock Germany Michael Page, Department of Chemical and Biological Sciences, The University of Huddersfield, Huddersfield, HD1 3DH, UK Francesca Paradisi, School of Chemistry and Chemical Biology, University College Dublin, Belfield, Dublin 4, Ireland Serge Parel, Biofocus DPI AG, Gewerbestrasse 16, CH-4123 Allschwil, Switzerland Teodor Parella, Servei de Ressona`ncia Magne`tica Nuclear, Universitat Auto`noma de Barcelona, 08193 Bellaterra, Barcelona, Spain Jaiwook Park, Department of Chemistry, Pohang University of Science and Technology (POSTECH), San-31, Hyojadong, Pohang 790-784, Korea Sara Pellegrino, Dipartimento di Scienze Molecolari Applicate ai Biosistemi (DISMAB), Facolta` di Farmacia, Universita` degli Studi di Milano, via Venezian 21, 20133 Milano, Italy

xxvi


Fabio Pezzotti, ECBB, ICMMO, Univ Paris-Sud, UMR 8182, F-91405 Orsay, France Robert S. Phillips, Department of Chemistry and of Biochemistry and Molecular Biology, University of Georgia, Athens, GA 30602, USA Umberto Piantini, Institute of Life Technologies, University of Applied Sciences Valais, Route du Rawyl 47, 1950 Sion, Switzerland David Pollard, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA Monique Ponelle, Analytical and Imaging Sciences, NIBR, Novartis Pharma AG, CH-4002 Basel, Switzerland Manfred T. Reetz, Max-Planck-Institut fu¨r Kohlenforschung, Kaiser-Wilhelm-Platz 1, 45470 Mu¨lheim/Ruhr, Germany Renate Reiss, School of Chemistry, Manchester Interdisciplinary Biocentre, University of Manchester, 131 Princess Street, Manchester, M1 7DN, UK Sergio Riva, Istituto di Chimica del Riconoscimento Molecolare, C.N.R., Via Mario Bianco 9, 20131 Milano, Italy Chris Roberge, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA J. Augusto R. Rodrigues, State University of Campinas, Institute of Chemistry, CP 6154, 13084-971, Campinas-SP, Brazil John P. N. Rosazza, Center for Biocatalysis and Bioprocessing, 2501 Crosspark Road, Suite C100 MTF, University of Iowa, Iowa City, Iowa, IA 52242-5000, USA Monika Rusek, Max-Planck-Institut fu¨r Kohlenforschung, Kaiser-Wilhelm-Platz 1, 45470 Mu¨lheim/Ruhr, Germany Floris P. J. T. Rutjes, Institute for Molecules and Materials, Radboud University Nijmegen, Toernooiveld 1, NL-6525 ED Nijmegen, The Netherlands Emmanuelle Sagot, Department of Chemistry, Universite´ Blaise Pascal, ClermontFerrand, France Francesca Sagui, Istituto di Chimica del Riconoscimento Molecolare, C.N.R., Via Mario Bianco 9, 20131 Milano, Italy Takashi Sakai, Division of Chemistry and Biochemistry, Graduate School of Natural Science and Technology, Okayama University, Tsushima, Okayama 700-8530, Japan Soth Samreth, Fournier Pharma, 50 rue de Dijon, 21121 Daix, France Christopher K. Savile, Department of Biochemistry, Molecular Biology & Biophysics and The Biotechnology Institute, University of Minnesota, 1479 Gortner Avenue, Saint Paul, MN 55108 USA Adrian Scaffidi, Chemistry M313, School of Biomedical, Biomolecular and Chemical Sciences, University of Western Australia, Crawley, WA 6009, Australia


xxvii

Andreas Schmid, Laboratory of Chemical Biotechnology, Faculty of Biochemical and Chemical Engineering, TU Dortmund, Emil-Figge-Strasse 66, 44221 Dortmund, Germany Jens Schrader, DECHEMA e.V., Karl-Winnacker-Institut, Biochemical Engineering Group, Theodor-Heuss-Allee 25, 60486 Frankfurt, Germany Frank Schulz, Max-Planck-Institut fu¨r Kohlenforschung, Kaiser-Wilhelm-Platz 1, 45470 Mu¨lheim/Ruhr, Germany Ioulia Smonou, Department of Chemistry, University of Crete, Iraklion-Voutes, 71003 Crete, Greece Timothy Snape, School of Chemistry, Manchester Interdisciplinary Biocentre, University of Manchester, 131 Princess Street, Manchester, M1 7DN, UK Georg A. Sprenger, Institute of Microbiology, University of Stuttgart, Allmandring 31, 70569 Stuttgart, Germany Robert V Stick, Chemistry M313, School of Biomedical, Biomolecular and Chemical Sciences, University of Western Australia, Crawley, WA 6009, Australia Matthew Stirling, Department of Chemical and Biological Sciences, The University of Huddersfield, Huddersfield, HD1 3DH, UK Takeshi Sugai, Faculty of Pharmacy, Keio University, 1-5-30, Shibakoen, Minato-ku, Tokyo 105-8512, Japan Peter Sutton, GlaxoSmithKline, Synthetic Chemistry, Gunnels Wood Road, Stevenage, Hertfordshire SG1 2NY, UK Andreas Taglieber, Max-Planck-Institut fu¨r Kohlenforschung, Kaiser-Wilhelm-Platz 1, 45470 Mu¨lheim/Ruhr, Germany Lixia Tang, Biochemical Laboratory, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Nijenborgh 4, 9747 AG, Groningen, The Netherlands Alan Taylor, Centre for Material Science, University of Central Lancashire, Preston Lancashire, UK David Tellers, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA Helene Therisod, ECBB, ICMMO, Univ Paris-Sud, UMR 8182, F-91405 Orsay, France Michel Therisod, ECBB, ICMMO, Univ Paris-Sud, UMR 8182, F-91405 Orsay, France Ai-Ming Tong, Laboratory of Biocatalysis and Bioprocessing, State Key Laboratory of Bioreactor Engineering, East China University of Science and Technology, Shanghai 200237, PR China Rafael de la Torre, Pharmacology Research Unit, Institut Municipal d’Investigacio´ Me´dica (IMIM), Barcelona, Spain

xxviii


Matthew Truppo, Process Research, Merck Research Laboratories, Merck & Co. Inc. Rahway, NJ, USA Nicholas J. Turner, School of Chemistry, Manchester Interdisciplinary Biocentre, University of Manchester, 131 Princess Street, Manchester, M1 7DN, UK Cleber V. Ursini, State University of Campinas, Institute of Chemistry, CP 6154, 13084971, Campinas-SP, Brazil Takamitsu Utsukihara, Department of Chemistry, Rikkyo (St.Paul’s) University, NishiIkebukuro, Toshima-Ku, Tokyo 171-8501, Japan J.A.J.M. Vekemans, Eindhoven University of Technology, Laboratory of Macromolecular and Organic Chemistry, PO Box 513, 5600 MB Eindhoven, The Netherlands Claire Vieille, Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824, USA Shaozhao Wang, Biotechnology Research Institute, National Research Council Canada, Montreal, Quebec H4P 2R2, Canada Marloes A. Wijdeven, Institute for Molecules and Materials, Radboud University Nijmegen, Toernooiveld 1, NL-6525 ED Nijmegen, The Netherlands Chi-Huey Wong, Department of Chemistry, The Scripps Research Institute, 10550 North Torrey Pines Rd., La Jolla, CA 92307, USA Matthias Wu¨st, Institute of Life Technologies, University of Applied Sciences Valais, Route du Rawyl 47, 1950 Sion, Switzerland Li Xu, Key Laboratory for Molecular Enzymology and Engineering of Ministry of Education, Jilin University, Changchun, 130021, PR China Jian-He Xu, Laboratory of Biocatalysis and Bioprocessing, State Key Laboratory of Bioreactor Engineering, East China University of Science and Technology, Shanghai 200237, PR China Hitomi Yamaguchi, Research & Development Center, Nagase & Co., Ltd., 2-2-3, Murotani, Nishi-ku, Kobe 651-2241, Japan Jianzhong Yang, Biotechnology Research Institute, National Research Council Canada, Montreal, Quebec H4P 2R2, Canada Toyokazu Yoshida, Department of Biomolecular Science, Gifu University, Yanagido 1-1, Gifu 501-1193, Japan Hui-Lei Yu, Laboratory of Biocatalysis and Bioprocessing, State Key Laboratory of Bioreactor Engineering, East China University of Science and Technology, Shanghai 200237, PR China Sheng Yuan, Nanjing Engineering Research Center for microbiology, Jiangsu Key Laboratory for Biodiversity and Biotechnology, College of Life Science, Nanjing Normal University, 1, Wenyuan Rd, Nanjing 210046, PR China


xxix

Francesca Zambianchi, Istituto di Chimica del Riconoscimento Molecolare, CNR, via Mario Bianco 9, 20131 Milano, Italy J. Gregory Zeikus, Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824, USA Karla I. Ziegelmann-Fjeld, Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824, USA Xue-Zhong Zhang, Key Laboratory for Molecular Enzymology and Engineering of Ministry of Education, Jilin University, Changchun, 130021, PR China Yongzhong Zhu, Department of Chemistry, National University of Singapore, Kent Ridge, Singapore 119260, Republic of Singapore

1 Biotransformations in Small-molecule Pharmaceutical Development Joseph P. Adams, Andrew J. Collis, Richard K. Henderson and Peter W. Sutton

1.1

Introduction

The demand for medicines that treat illnesses formerly associated with the developed world is expanding at a time when some countries are becoming increasingly affluent. As a result, the global pharmaceutical market is predicted to grow to $800 billion by the year 2020.1 However, as demand increases for products, the pharmaceutical industry is facing increasing pressures that can primarily be attributed to three factors: 1. As the global population ages and lifestyles become more sedentary, the cost of healthcare is becoming increasingly unsustainable. This is no more so than in the USA, where, although prescription products contribute only 10 % of healthcare costs, they are perceived to be much higher by the consumer and so represent an easy political target for cost cuts through price controls. 2. Erosion of product lifetimes as a result of greater generic competition means that a product can expect to lose the majority of its market in as little as 3 months after patent expiry. 3. Spiralling R&D costs. Typically, it takes 10 years at a cost of $500 million to bring a drug to market.2 Fewer new molecular entities (NMEs) and biologics are reaching the market as a result of a shift of research focus away from already established and crowded therapeutic areas into new, unproven biological areas (Figure 1.1).3

Practical Methods for Biocatalysis and Biotransformations Ó 2009 John Wiley & Sons, Ltd

Edited by John Whittall and Peter Sutton

Biotransformations in Small-molecule Pharmaceutical Development 60

R&D Spending (US$ Millions)

50000 45000

50

40000 35000

40

30000 30

25000 20000

20

15000 10000

10

5000

No. of NMEs and Biologics Approved

2

0

19 95 19 96 19 97 19 98 19 99 20 00 20 01 20 02 20 03 20 04 20 05 20 06

0

R&D Spending

NMEs and New Biologics Approved

Figure 1.1 R&D spending versus the number of NMEs and biologics approved by the US Food and Drug Administration (FDA). (Reprinted with permission from Pharma 2020: The vision: Which path will you take?, PricewaterhouseCoopers, 2007.)

NCEs

20 18 16 14 12 10 8 6 4

20 03

20 02

20 01

20 00

19 99

19 98

19 97

19 96

19 95

19 94

19 93

19 92

2 0

Year of Launch

achiral

chiral

Figure 1.2 Number of chiral and achiral marketed NCEs. (Reprinted with permission from Farina, V., Reeves, J.T., Senanayake, C.H. and Song, J.J. Asymmetric synthesis of active pharmaceutical ingredients. Chem. Rev. 2006, 106, 2734–2793. Copyright 2006, American Chemical Society)

Whereas new drugs reaching the market do not necessarily look any more complex or contain any more stereocentres than in the past (Figure 1.2),3 the complexity of drug candidates under development has increased on average. In addition, following the FDA’s 1992 policy statement on stereoisomers, it is now clearly more economical to progress an active pharmaceutical ingredient (API) in enantiopure form, as can be seen from the trend towards the launch of single-enantiomer new chemical entities (NCEs) (Figure 1.3).

1.2 Current Status of Biocatalysis

3

14 12

NCEs

10 8 6 4 2

20 03

20 02

20 01

20 00

19 99

19 98

19 97

19 96

19 95

19 94

19 93

19 92

0

Year of Launch

single enantiomer

racemate

Figure 1.3 Number of single enantiomer versus racemic NCEs. (Reprinted with permission from Farina, V., Reeves, J.T., Senanayake, C.H. and Song, J.J. Asymmetric synthesis of active pharmaceutical ingredients. Chem. Rev. 2006, 106, 2734–2793. Copyright 2006, American Chemical Society)

This trend seems likely to continue, with over 50 % of current drug candidates being developed as single enantiomers.4 An obvious consequence has been an explosion of research activity into asymmetric synthetic methods. These combined issues, on average, make pharmaceutical companies as much as 50 % riskier than other big industries.5 Led by the FDA’s Current Good Manufacturing Practices for the 21st Century initiative,6 the pharmaceutical industry has begun to apply a risk management and quality systems approach, practiced in some other industries for decades, to products throughout their lifetimes.7 The guidance from the FDA’s Process Analytical Technologies regulatory framework, developed over the last decade, aims to build quality by design into pharmaceutical products through better process understanding and increased innovation. The ultimate goal is to minimize risk to the patient whilst encouraging the industry to cut operating costs.8 Encouraged by this more flexible regulatory approach, there is an increased willingness within the industry to adopt ‘new’ technologies.9

1.2

Current Status of Biocatalysis

A biotransformation, as defined by Straathof et al.,10 is ‘a process that describes a reaction or a set of simultaneous reactions in which a pre-formed precursor molecule is converted using enzymes and/or whole cells, or combinations thereof, either free or immobilised’. Fermentation processes, with de novo product formation from a carbon and energy source, such as glucose via primary metabolism, are outside the scope of this chapter and book unless employed in conjunction with a biotransformation. Biocatalysis has long been known as a green technology, capable of delivering highly stereo-, chemo- and regioselective transformations that can sometimes allow

4

Biotransformations in Small-molecule Pharmaceutical Development Polymers Cosmetics Food Animal Feed Agro Other sectors Pharma

Figure 1.4 Number of biotransformations used catagorised by industrial sector (based on 134 processes). (Reprinted from Straathof, A.J.J., Panke, S. and Schmid, A. The production of fine chemicals by biotransformations. Curr. Opin. Biotechnol. 2002, 13, 548–556 with permission from Elsevier.)

the number of steps in a synthetic route to be reduced. Numerous industrial biotransformations (announced to be commercialized at a scale of >100 kg per annum) are in operation worldwide, many of which have been described by Liese et al.11 Most of these known biotransformations are used to produce building blocks that are subsequently supplied to the pharmaceutical industry (Figure 1.4).10 Biocatalysis is still an emerging field; hence, some transformations are more established than others.12 Panke et al.13 have performed a survey of patent applications in the area of biocatalysis granted between the years 2000 and 2004. They found that although hydrolases, which perform hydrolyses and esterifications, still command widespread attention and remain the most utilized class of enzyme (Figure 1.5), significant focus has turned towards the use of biocatalysts with different activities and in particular alcohol dehydrogenases (ADHs) – also known as ketoreductases (KREDs) – used for asymmetric ketone reduction. Whereas the number of industrial biotransformations ‘known’ to be operating in 2002 was 134, the number of chiral drug candidates is much greater. Farina et al.3 have estimated between 500 and 1000 single-enantiomer APIs to be in development each year in the global pipeline. This implies that biotransformations might supply only a small percentage of chiral centres. This might be partially attributable to the reluctance of the pharmaceutical industry to innovate in the absence of the recently established regulatory directives, or to the lack of commercial enzymes available on a large scale. However, the main factor lies in the strategy used to incorporate chirality into drug candidates. Oxidoreductases Oxidising cells Reducing cells Isomerases Lyases Hydrolases Transferases

Figure 1.5 Enzyme Types Used in Industrial Biotransformations (based on 134 processes). (Reprinted from Straathof, A.J.J., Panke, S. and Schmid, A. The production of fine chemicals by biotransformations. Curr. Opin. Biotechnol. 2002, 13, 548–556 with permission from Elsevier.)

1.2 Current Status of Biocatalysis

5

In line with the construction of a target molecule from smaller, complex fragments, it is generally preferred to introduce chirality into a synthetic route at an early stage through the purchase of simple chiral starting materials from the fine chemical industry. This has been demonstrated by Carey et al.,14 who performed a survey of 128 drug candidate syntheses, many of which were in an early phase of development. They found that, of the 69 chiral drug candidates considered, 55 % of the 135 chiral centres present were bought in from the fine chemicals industry. In cases where it was necessary to generate chirality in-house, the favoured method was racemate resolution (28 % of chiral centres – with classical salt formation employed in two-thirds of cases and dynamic kinetic resolution, chromatography and biocatalytic methods evenly distributed in the remainder) followed by chemical asymmetrization (10 % of chiral centres – see Section 1.3.4 for definition) and diastereoselective induction (7 % of chiral centres). Another important source of chirality (which was not exemplified in that article) is fermentation technology, which provides access to many of the important chiral scaffolds that have been employed by the industry in wellknown classes of drug, such as b-lactam antibiotics and, more recently, first-generation statins. Some 35 % of the chiral building blocks that are bought in from the fine chemical industry, such as both proteinogenic and non-proteinogenic amino acids, carboxylic acids, amines, alcohols and epoxides, are produced using generic biocatalytic technologies, and this is expected to increase to 70 % by 2010.12 Far more chiral centres present in APIs are derived from industrial biotransformations than would be expected by counting the number of known processes.15 This can also be noted from the procedures given in this book, the vast majority of which provide biocatalytic routes to chiral building blocks. These chiral building blocks, in turn, will be dictated by current drug candidates within the pharmaceutical industry’s pipeline. Given the wide utility of biocatalysis in the fine chemical industry, why is there such an in-house reliance on classical methods of enantioseparation? In fact, why is biocatalysis not applied more generally as a replacement for atom-inefficient or hazardous reactions that are intensively used in the pharmaceutical industry, such as amidation, reduction and oxidation?16 The sparse incorporation of biocatalysts into the process chemist’s toolbox is at least in part due to a number of long-standing issues that differ depending on the drug development phase. At an early stage of development, where little resource is available for new route development, biocatalysis options are often neglected due to a lack of sufficient commercially available biocatalysts.17 In contrast, classical salt formation regularly provides access to chiral material in >99.5 % enantiomeric purity; hence its widespread adoption. At a later phase biocatalysis may be considered, but the longer development times often needed and the more advanced state of competing chemical routes put it at a disadvantage. Many of these issues have been or are being addressed. For example, with the continued expansion in the number of microorganisms whose genomes have been sequenced, the application of bioinformatics techniques is leading to a rapid expansion in the number of commercially available enzymes such as ADHs (ketone reduction), nitrilases (nitrile hydrolysis), enoate reductases (,b-unsaturated olefin reduction) and transaminases (reductive amination).18 Having identified a putative enzyme gene by sequence similarity, it can now be quickly and cheaply generated by using oligonucleotide synthesis services that are provided by a number of companies. However, it is predicted that about 99 % of

6

Biotransformations in Small-molecule Pharmaceutical Development

microorganisms are ‘non-cultivable’, and metagenomics – the extraction of environmental DNA – is proving highly successful in accessing novel biocatalysts from this untapped resource.19 Further expansion in the number of commercially available enzymes and the modification of hits to suit process requirements is being fuelled by advances in enzyme engineering20 and high-throughput screening (HTS) technologies.21 A particularly elegant approach is the protein sequence activity relationship (ProSAR) technology developed by Codexis.22 By using a multivariate analysis approach, libraries of enzyme variants containing programmed mutations generated from different sources of diversity are screened against a given substrate and sequenced. Positive mutations and interactions between different mutations can then be ascertained, allowing more active variants to be predicted in silico, thus reducing the bottleneck often caused by screening. As the impact of individual mutations is understood, the possibility of missing important ones or carrying false hits through to the next round is reduced compared with traditional hit-based directed-evolution strategies. Similar multivariate techniques are used on a daily basis within the pharmaceutical industry (quantitative structure–activity relationships in medicinal chemistry and design of experiments to understand and optimize chemical reactions during process development) to interpret complex data sets. So powerful is this technique at producing ADHs suitable for process applications that one pharmaceutical company now considers biocatalysis as their first option for asymmetric ketone reduction. There is also a greater appreciation within the biotech community that alternative routes to a drug candidate are always available and, although they are sometimes technologically inferior, will always be favoured if freedom to operate is at stake.

1.3

Application of Biocatalysis in the Pharmaceutical Industry

This section primarily focuses on examples of biotransformations that have been developed for the preparation of small-molecule APIs (molecular weight 80 % isolated yield by the treatment of ribavirin with 0.8 weight equivalents of CALB in tetrahydrofuran (THF) at 60 °C for 24 h. This approach has also been used to produce ester prodrugs of other nucleoside antivirals, such as nelarabine.70 H2NOC

H2NOC

N N

N

CALB, acetone oxime,

O

HO HO

OH

CbzNH

CO2H

O CbzHN

N N

N

O

O HO

OH

Ribavirin

Scheme 1.19

Regioselective preparation of 50 -N-CBz-(S)-alaninyl ribavarin

The preparation of N-CBz-(S)-valinyl lobucavir provides a particularly challenging example, where only one of two primary alcohols is acylated with excellent regioselectivity (Scheme 1.20).71

1.3 Application of Biocatalysis in the Pharmaceutical Industry O N N

O N

NH N

HO

25

NH2

NH

N

Immobilized PCL,

N

HO O2N

O

O

OH

NH2

O

O

NHCbz

NHCbz

Scheme 1.20 Regioselective esterification of lobucavir (PCL: Pseudomonas cepacia lipase, now known as Burkholderia cepacia) O

HO O H

O O

Lovastatin

O

HO O H

O O

Simvastatin

Figure 1.6 Structures of lovastatin and simvastatin.

Simvastatin is a semi-synthetic statin that is produced from the natural statin lovastatin.72 Both are potent antihypercholesterolemic agents with simvastatin differing from lovastatin by just one additional methyl substituent residing on the 2-(S)-methylbutyrate side chain (Figure 1.6). Lovastatin is produced by fermentation from the filamentous fungus Aspergillus terreus and can be converted to simvastatin by a single-step chemical methylation.73 However, this transformation is hampered by low yields, which result in downstream processing issues resulting from difficulties in the separation of starting material and product. Simvastatin is instead produced using a lengthier protection/deprotection strategy.74 To overcome these separation issues, Schimmel et al.75 sought a hydrolase enzyme capable of selectively hydrolysing the 2-(S)-methylbutyrate ester of lovastatin ammonium salt (LAS) whilst leaving the more hindered 2-dimethylbutyrate ester of the simvastatin ammonium salt (SAS) unchanged. After screening 150 microorganisms, the fungus Clonostachys compactiuscula was found to produce a suitable esterase. By applying this esterase to inseparable LAS/SAS mixtures resulting from the single-step chemical methylation, they were able to hydrolyse LAS selectively to the more polar, readily separable monacolin J ammonium salt, thus providing a two-step conversion of lovastatin to simvastatin (Scheme 1.21). Regioselective esterification of the 8-hydroxyl group of accumulated monacolin J, produced using a truncated lovastatin biosynthetic pathway, could provide a viable biocatalytic route to simvastatin. With this aim, Xie and Tang76 cloned and expressed the acyl transferase LovD from the lovastatin biosynthetic pathway into E. coli; they found

26

Biotransformations in Small-molecule Pharmaceutical Development HO

HO

CO2NH4 OH

O

O

O

HO

CO2NH4 OH

CO2NH4 OH

OH

O Esterase from

+

+

SAS

C. compactiuscula LAS

Monacolin J ammonium salt

SAS

Scheme 1.21 Selective enzymatic hydrolysis of LAS/SAS mixtures

that LovD was not only active towards lovastatin synthesis, but also capable of producing simvastatin using simple -dimethylbutyrate thioesters (Scheme 1.22). Whereas a biotransformation using partially purified LovD in aqueous solution gave a conversion of only 60 % due to competing hydrolysis, whole-cell reactions went to completion to afford 4–6 g L1 product concentrations. The authors speculated that the superior results obtained from the whole-cell reactions might result from active transport of simvastatin out of the cells which are subsequently impermeable to re-entry, whereas the more polar monacolin J can diffuse in both directions. The efficiency of the transformation was later improved by knocking out the gene expressing the BioH enzyme which is responsible for competing thioester hydrolysis.77

O H OH

O

HO

O

HO

O H

O LovD

O

O S Monacolin J

Simvastatin

Scheme 1.22 Biocatalytic regioselective acylation of monacolin J

Using molecular biology techniques to redirect primary metabolic pathways, microorganisms may be engineered to overproduce a wide range of biochemical intermediates, such as amino acids and vitamins.78 This principle can be extended by introducing novel enzymes and, thereby, novel biotransformation steps into microbial hosts in order to generate unnatural products from natural precursors. Such a modification may be lethal for the host cell, requiring the application of techniques developed for controlled, conditional gene expression in the production of recombinant proteins.79 This is illustrated by the engineered microbial production of the nucleoside thymidine (TdR), an important starting material for synthesis of the antiretrovirals zidovudine and stavudine (Scheme 1.23). Although thymidine-50 -triphosphate is an almost universal component of DNA, it is exclusively derived from thymidine-50 -monophosphate (TMP). In contrast, TdR does not occur naturally and so it is impossible to manufacture TdR by manipulation of existing metabolic pathways as for most biochemical intermediates. This problem was addressed

1.3 Application of Biocatalysis in the Pharmaceutical Industry

27

O NH N O

O

N O

O

O

O

NH

NH O HO P O OH

HO

N

Phosphohydrolase

OH Thymidine-5'-monophosphate (TMP)

HO

O

Stavudine O O

OH

NH

Thymidine (TdR)

N HO

O

O

N3 Zidovudine

Scheme 1.23 Enzyme-catalysed hydrolysis of thymidine-50 -monophosphate

by McCandliss and Anderson80 using a gene encoding TMPase, a phosphohydrolase acting on TMP to generate TdR. Such enzymes are found only in rare bacterial viruses with DNA incorporating uracil in place of thymine. This potentially lethal gene, capable of knocking out normal DNA synthesis, was coupled to an inducible promoter allowing strict control of its expression. Typical genetic refinements used in metabolic engineering were introduced, knocking out enzymes that would degrade TdR and enhancing pathways leading into TMP synthesis. Deoxyribonucleotides are derived metabolically by reduction of the corresponding ribonucleotide, an arrangement that reflects the greater abundance of RNA compared with DNA. This reduction occurs at the level of the nucleoside diphosphates. TMP is derived by methylation of 20 -deoxyuridine-50 -monophosphate (dUMP), itself derived from the corresponding 20 -deoxyuridine-50 -diphosphate, which is generated by reduction of the analogous ribonucleotide uridine-50 -diphosphate. In order to enhance the process to commercial levels of productivity in an engineered strain of E. coli, Anderson et al.81 added a number of recombinant genes encoding both a ribonucleotide reductase and the thioredoxin required to regenerate its reduced and active form. TMPase acts to dephosphorylate both TMP and its precursor dUMP, forming a mixture of TdR and 20 -deoxyuridine (UdR). As a starting material for zidovudine synthesis, TdR must be essentially free of this impurity, which would pass through the manufacturing process to form a demethylated analogue of zidovudine. Separation of TdR and UdR requires difficult and costly downstream processing; hence, the key to a commercial process is metabolic engineering to minimize biosynthetic UdR. Anderson et al.82 investigated a range of solutions to this problem, each based on the principle of efficient methylation of dUMP to TMP to avoid the accumulation of a significant pool of free dUMP that could be converted to UdR. Various techniques were used to increase the efficiency of the methylation reaction itself using alternative forms of thymidylate synthase with enhanced catalytic activity and altered regulation. However, the most significant improvement was by enhancing activity of the enzymes recycling and replenishing the methyl donor 5,10-methylenetetrahydrofolate (Scheme 1.24).

28

Biotransformations in Small-molecule Pharmaceutical Development +

NADP (regeneration)

THFo

dihydrofolate reductase (EC 1.5.1.3) +

NADPH + H

CH2-THFo

DHFo thymidylate synthase (EC 2.1.1.145)

O

O

NH O HO P O OH

N O

NH

O

O HO P O OH

OH

N O

O

OH

dUMP

TMP

thymidylate synthase (EC 2.1.1.148)

FADH2 CH2-THFo

FAD THFo CH2-THFo THFo DHFo NADPH + NADP FADH2 FAD

5,10-methylenetetrahydrofolate tetrahydrofolate dihydrofolate reduced nicotinamide adenine dinucleotide phosphate nicotinamide adenine dinucleotide phosphate reduced flavin adenine dinucleotide flavin adenine dinucleotide

(regeneration)

Scheme 1.24 Methylation in TMP biosynthesis

Biosynthetic production of thymidine is overall a complex process combining the controlled introduction of a novel biotransformation step into a biological system with selective enhancement or knock-out of a series of existing metabolic steps. Metabolic engineering to enhance cofactor recycling at both ribonucleotide reduction and dUMP methylation steps has important parallels in other systems, as whole-cell biotransformations are frequently employed as a means to supply, in situ, high-cost and usually labile cofactors. Atorvastatin, an antihypercholesterolemic agent, is a synthetic drug that was initially produced in kilogram quantities using an 11-step chemical route. The syn-1,3-diol-containing side chain was produced from the chiral starting material, isoascorbic acid (Scheme 1.25).83 Numerous biocatalytic routes to this challenging intermediate have been reported.84 For example, Fox et al.85 have recently developed an efficient regioselective cyanation starting from low-cost epichlorohydrin (Scheme 1.26). Initial experiments found that halohydrin dehydrogenase from Agrobacterium radiobacter expressed in E. coli produced the desired product, but inefficiently. To meet the projected cost requirements for economic viability, the product needed to be produced at 100 g L1 with complete conversion and a 4000-fold increase in volumetric productivity. The biocatalyst needed to function under neutral conditions to avoid by-product formation, which causes downstream processing issues. Using ProSAR technology (see Section 1.2), the group identified a variant halohydrin dehalogenase containing 37 mutations that gave the required volumetric productivity increase. This methodology is also applicable to other antihypercholesterolemic drugs, such as rosuvastatin and fluvastatin (Figure 1.7).


29

OH HO

OH

O O

i) TBDMSCl, Im, DMAP CO2Me

Br

ii) NaCN, DMSO

HO

OTBDMS CO2H

NC

iii) NaOH

OH

OH O

i) CDI, Mg(O2CCH2CO2tBu)2 ii) Bu4NF, AcOH, THF

O

o

CO2tBu

NC

i) NaBH4, Et2BOMe, MeOH, –90 C

O

NC

CO2tBu

ii) Me2C(OMe)2, MeSO3H

OH

O O

H2, Raney Ni, MeOH, 50 psi

O

CO2H

N H

CO2tBu

H2N

OH

N

F Atorvastatin

Scheme 1.25 Chemical synthesis of the atorvastatin side chain

OH Cl

OH

Halohydrin dehalogenase CO2Et

NaCN

O

Scheme 1.26

CO2Et

NC

CO2Et

Halohydrin-catalysed cyanation of epichlorohydrin

F S O

F

N

N O N

CO2H OH

Rosuvastatin

OH

CO2H

N OH OH Fluvastatin

Figure 1.7 Some other statins containing a 1,3-syn-diol side chain.

In conclusion, regioselective biocatalysis has been extensively employed to access both semi-synthetic and synthetic pharmaceuticals. The methodology is particularly attractive for the streamlining of processes through the elimination of protecting-group strategies and to avoid the use of hazardous reagents.

30

1.3.3


Diastereoselective Biotransformations

Diastereoselective reactions, where one or more chiral centres are generated in a selective manner within a molecule that already contains chirality, to produce single diastereoisomers (epimers) are very common in nature. Some examples of chemical processes which harness the properties of biocatalysts are shown below. Highly diastereoselective enzyme-catalysed glycosylation reactions allow access to functionalized sugars and highly complex polysaccharides and provide an important pathway by which xenobiotics are metabolized (see Section 1.3.1). A similar transformation is the nucleoside phosphorylase (NP)-catalysed reversible cleavage of the N-glycosidic linkage of a nucleoside in the presence of phosphate to generate the corresponding pentose sugar phosphate and free nucleobase. NPs are ubiquitous in biology, and substrate ranges include deoxyribonucleosides and/or ribonucleosides with purine or pyrimidine nucleobases. N-Transglycosylation can be achieved by coupling cleavage of the sugar from a donor nucleoside to synthesis of a new nucleoside using a second, acceptor base. This reaction, which is completely regioselective towards the base and diastereoselective towards formation of the b-anomer at the sugar is a useful strategy for synthesis of nucleoside analogues, including many antiviral and anticancer agents, such as ribavirin or, indirectly via thymidine, zidovudine and stavudine (Scheme 1.27).86

N

HN H2N

O HO

HN

HN

N

N

HO

O

O

O

O HO

OH

HO

Guanosine

O

N O

N O

HO HO

OH

Thymidine

Methyluridine

O

O H2NOC

HO

O

N

O

HO

HN

HN

N N

HO

Scheme 1.27

N O

HO N3

OH

Ribavirin

O

N O

Stavudine

Zidovudine

Antiretroviral nucleosides accessible by NP catalysis

Using guanosine or 20 -deoxyguanosine as starting material for the synthesis of ribonucleosides or deoxyribonucleosides respectively, the reaction can be driven towards completion by precipitation of the highly insoluble guanine co-product. This approach has

1.3 Application of Biocatalysis in the Pharmaceutical Industry O

H2N

N N H

+

N

N O

HO HO

H2NOC

CONH2

N

HN

N

N

O

N N

H3PO4, bacterial cells, 60 °C

Guanosine

N

N H

OH Guanine (precipitated)

Ribavirin

Scheme 1.28

N

HN H2N

HO

OH

+

O

HO

31

Enzymatic direct synthesis of ribavirin

been used for direct synthesis of the antiviral ribavirin in approximately 75 % yield using bacterial cells (Brevibacterium) (Scheme 1.28).87 Like many reported N-transglycosylations, this reaction uses uncharacterized nucleoside phosphorylases from whole cells held at 50–60 °C, a temperature well above the range for viability of the parent microorganism. Remarkable temperature stability has been reported for three well-known NPs of E. coli: purine nucleoside phosphorylase (PNP), uridine phosphorylase (URDP) and thymidine phosphorylase.88 Certain NPs can use pentoses other than ribose or deoxyribose as substrates, enabling the synthesis of nucleoside analogues with unnatural sugar moieties: for example, in the synthesis of purine arabinonucleosides.89 A convenient donor for transarabinosylation is 9-b-D-arabinofuranosyluridine (Ara-U), which can be accessed from uridine using a twostep chemical process to invert the 20 -hydroxyl group.90 A general protocol for preparation of purine analogues using Ara-U with a mixture of purified URDP and PNP from E. coli is described by Averett et al.95 The enzymes are used in varying proportions, depending upon the reaction rate for the required purine nucleoside synthesis, and are sufficiently robust for addition of water-miscible solvents to aid substrate solubility. The URDP/PNP/Ara-U process is used to manufacture nelarabine, a water-soluble prodrug of 9-b-D-arabinofuranosylguanidine produced as a treatment for acute lymphoblastic leukaemia (Scheme 1.29).70,92 The two-enzyme process is run at 200 g L1 O HN O O

N O

HO

+

URDP

H3PO4

HO

HO

HN O

OH

O

OPO3H2

+ N H

HO

OH

α-D-Arabinose-1-phosphate

Ara-U

OMe OCH3 N

N H2N

+ N

N H

O

HO

N

N

OPO3H2 PNP

H 2N

HO

+

O

HO

6-Methoxyguanine

N

N

OH HO

OH

Nelarabine

Scheme 1.29

Preparation of nelarabine from Ara-U

H3PO4

32


substrate concentration and can be driven to 90 % conversion over 50 h by using the correct ratio of the two enzymes. As with other NP-catalysed transformations, the process is run at 50 °C. To improve thermostability and facilitate reuse, the enzymes are co-immobilized onto the same support. For design of a simple manufacturing process, the thermostability of the NP enzymes is a very useful feature. Although heat treatment can be used as part of a purification protocol to isolate the enzymes from contaminating materials, the high temperature of operation itself excludes undesired enzyme-catalysed side reactions. For example, in the synthesis of 9-b-D-arabinofuranosyladenine from Ara-U and adenine, using a wet cell paste of Enterobacter aerogenes, adenine and Ara-U mainly underwent deamination at lower temperatures to form hypoxanthine and uracil respectively.93 At elevated temperature, deamination was completely eliminated and the rate of transarabinosylation increased. One drawback of biocatalysis is that enzymes are not available in both enantiomeric forms. Particularly where a class of enzymes whose natural substrates are optically active, such as nucleosides, it can be difficult if not impossible to find an alternative enzyme that will accept the unnatural substrate enantiomer. This is not insurmountable if directedevolution approaches are used, but it can be prohibitively expensive, especially when the desired product is in an early stage of development or required for use only as an analytical reference or standard. During the development of nelarabine, the opposite enantiomer (ent-nelarabine) was required as an analytical marker.94 The lengthy chemical route to ent-nelarabine and the fact that this chemical route is necessary illustrate both the advantages and disadvantages of biocatalytic approaches. The chemical synthesis of ent-nelarabine is not straightforward, commencing with a three-step global acetylation of the sugar (Scheme 1.30). As chemical glycosylation using arabinose results predominantly in formation of the undesired -anomer, ribose is instead used as the starting sugar. The enhanced diastereoselectivity HO HO

O

OH

O

MeO

MeOH, HCl

OH

HO

O

MeO

Ac2O, pyridine

OH OH

AcO

OAc OAc

Cl AcOH, Ac2O, H2SO4

AcO

OAc

BSA, TMSOTf, MeCN, 75 °C H2N

AcO

N H

H2N

N

OAc OAc

NH2 OMe

N N

O

AcO

N

OMe N

N

N

Cl

OAc N

NaOMe, MeOH

N

N

O

N

HO

O

OH

N

N

5 steps H2N

OH

Scheme 1.30 Chemical synthesis of ent-nelarabine

N

N

HO

O

OH OH


33

gained by the use of ribose in the glycosylation reaction also has a penalty, in that five additional steps are required in order to invert the 20 -alcohol of the resultant b-riboside. Furthermore, the unnatural 6-methoxyguanine base reacts chemically at N-7, as opposed to N-9 selectivity of the biocatalytic approach. This could be rectified by instead using 6-chloroguanine (to deactivate N-7) which could later be converted to the methoxide with concomitant acetate deprotection. Thus, ent-nelarabine was produced using an overall 10-step procedure. Carbon–carbon bond lyases, used in the reverse, synthetic direction have also enjoyed significant application in the pharmaceutical industry. For example N-acetyl-D-neuraminic acid (NANA), an intermediate in the chemoenzymatic synthesis of the influenza virus sialidase inhibitor zanamavir, may be synthesized using NANA aldolase. In nature, NANA arises through condensation of phosphoenolpyruvic acid with N-acetyl-D-mannosamine (NAM) catalysed by the biosynthetic enzyme NANA synthase.95 Owing to the labile nature of phosphoenolpyruvate, the use of this reaction in the synthesis of NANA has been limited to whole-cell systems where this substance can be generated biosynthetically in situ.96 Most recently, the NANA synthase reaction forms the basis of fermentation processes for total biosynthesis of NANA.97 Catabolic enzyme NANA aldolase catalyses cleavage of NANA to form NAM and pyruvic acid, the latter being a more attractive material for a chemoenzymatic process. It has long been known that the reverse reaction may be used for NANA synthesis.98 However, this approach to a manufacturing process also has complications. NAM is produced by base-catalysed epimerization of N-acetyl-D-glucosamine (NAG), generating an unfavourable 1:4 mixture of NAM:NAG. NAG, although not a substrate for the aldolase, inhibits the reaction. In addition, excess pyruvate is required to push the equilibrium in favour of product formation (Scheme 1.31). Although 90 % yields can be obtained at laboratory scale using E. coli NANA aldolase using a NAG:NAM mixture, the NANA product is difficult to separate from the excess pyruvate required to achieve this.

O

OH NHAc

HO

O

epimerization

HO

OH

HO

(base or enzyme catalysed)

NHAc

HO

OH

OH

NAG

NAM HO

immobilized NANA-aldolase

OH O

HO (excess) CO2H

OH CO2H

O

OH

HO

O HO

CO2H

AcHN

AcHN OH NANA

HN

NH2 NH

Zanamavir

Scheme 1.31

Aldolase-catalysed preparation of NANA

34


Cipolletti et al.99 recently described a crystallization procedure to isolate NAM in >98 % purity from a 4:1 NAG:NAM epimerate, potentially enabling the use of a NAG-free process. However, Mahmoudian et al.100 developed a scalable process using selective precipitation of NAG from aqueous solutions of NAG/NAM epimerate with isopropanol to generate a NAM-enriched solution as substrate for the enzymatic synthesis. Precipitated NAG could be recycled by base-catalysed epimerization. The NAM-enriched starting material allowed NANA product concentrations of 155 g L1 to be attained by using just two equivalents of pyruvate. Because of the lower pyruvate content, NANA could be purified by a simple crystallization following removal of the Eupergit C-immobilized aldolase by filtration. As an alternative to chemical epimerization, NAG epimerase may be used to maintain a constant NAM:NAG ratio in a one-pot reaction with pyruvate and NANA aldolase.101 The epimerase is itself inhibited by pyruvate, which must, therefore, be added continuously or via aliquots to the reaction. In a refined version of this reaction at laboratory scale, Kragl et al.102 produced NANA by a continuous process, using a membrane reactor to contain both enzymes in solution. 1.3.4

Asymmetric Biocatalysis

Asymmetric synthesis can refer to any process which accesses homochiral products. We will focus on asymmetric synthesis from racemic or prochiral starting materials in the presence of an enantioselective catalyst (enzyme). There are four general methodologies commonly applied: kinetic resolution, dynamic kinetic resolution, deracemization and asymmetrization. The process of obtaining homochiral product from a racemate is known as kinetic resolution. Kinetic resolution functions by the transformation of two enantiomers of a racemic mixture at different rates. The objective is to effect a change in the physical properties of one enantiomer to such an extent that the resulting product is readily separable from the other. The technique suffers from the inherent inability to access >50 % of the desired enantiomer unless the unwanted enantiomer can be racemized and recycled or inverted. Dynamic kinetic resolution (DKR) is an extension to the kinetic resolution process, in which an enantioselective catalyst is usually used in tandem with a chemoselective catalyst. The chemoselective catalyst is used to racemize the starting material of the kinetic resolution process whilst leaving the product unchanged. As a consequence, the enantioselective catalyst is constantly supplied with fresh fast-reacting enantiomer so that the process can be driven to theoretical yields of up to 100 %. There are special cases where the starting material spontaneously racemizes under the reaction conditions and so a second catalyst is not required. An alternative method of obtaining theoretical yields of up to 100 % of homochiral product from racemic mixtures is known as deracemization. This process again employs two catalysts in tandem and so bears much similarity to the DKR process. However, here an enantioselective catalyst preferentially transforms one enantiomer of starting material into a prochiral product. The prochiral product is then converted back into racemic starting material using an achiral catalyst, resulting in an overall enrichment towards one enantiomer of starting material. Further enrichment results by allowing the process to run over multiple cycles, until only one enantiomer remains.

1.3 Application of Biocatalysis in the Pharmaceutical Industry D

D Desymmetrisation C

X

Pro-R

A

D X

C

B Pro-S

35

B

(R )-product

Re Asymmetric Transformation

D C

D Si

X B

A X

C

B

(S)-product

Where X = carbon or heteroatom A,B,C and D are any substituent in decreasing CIP priority

Figure 1.8 Schematic representation of asymmetrization reactions.103 (Reprinted with permission from the American Chemical Society Copyright (2005))

The process of obtaining homochiral product from a prochiral starting material is known as asymmetrization. This encompasses reactions where a faster rate of attack of a reactive species occurs on one enantiotopic face of a prochiral trigonal biplanar system, or at one enantiotopic substituent of a C2 symmetrical system, resulting in the preferential formation of one product enantiomer. The latter is also frequently referred to as the ‘meso-trick’ or ‘desymmetrization’. These transformations can be more easily defined in pictorial form (Figure 1.8). Unlike kinetic resolution, catalytic desymmetrization and asymmetrization can afford enantiopure products in theoretical yields of 100 % and are more generally applicable than DKR or deracemization techniques. This section will only discuss examples of catalytic kinetic resolution, DKR, desymmetrization and asymmetrization. Deracemization will not be considered because, although an important developing technology, examples of its application to the production of chiral late-stage intermediates in API production have yet to appear. 1.3.4.1

Kinetic Resolution

This technique can allow the rapid development of processes for the separation of large quantities of enantiomers and can be ideal for early-stage ‘fit for purpose’ campaigns (where little resource is allocated to process development) in spite of the limitation in attainable yield. This can be useful in providing sufficient homochiral product for biological evaluation and the preparation of analytical standards of both enantiomeric forms. Most kinetic resolutions of pharmaceutical intermediates that have been reported involve the use of hydrolases, particularly lipases and proteases. This is because many hydrolases are commercially available (in bulk and kit form),104 do not require cofactors and are active in many organic solvents (see Section 1.4). Processes can therefore, often be developed rapidly, using high substrate concentrations and without specialist knowledge.

36


A second-generation manufacturing process involving a highly enantio- and diastereoselective lipase-catalysed kinetic resolution step has recently been reported for the production of pregabalin, a lipophilic g-aminobutyric acid analogue that was developed for the treatment of several central nervous system disorders (Scheme 1.32).105 CO2Et CO2Et CO2Et

CN Lipolase

CO2Et CN

pH 7, 24 h

CO2H

+

NH2

CO2Et CO2H

Pregabalin

CN

Scheme 1.32

Kinetic resolution of a key intermediate to pregabalin

Following a screen of hydrolase enzymes, the lipase from Thermomyces lanuginosus was selected based on its high activity and enantioselectivity. This enzyme is commercially available in industrial quantities as Lipolase, a cheap catalyst of importance to the detergents industry due to its high thermal stability and broad pH tolerance. Product inhibition was observed at concentrations over 1 M, and so divalent ion species were added as complexation agents. In the presence of calcium acetate, the reaction proceeded to completion at substrate concentrations up to 3 M, although only substoichiometric quantities were required, implying that the additive plays a more complex role than envisaged from the original rationale. A high concentration resulted in the added benefit of dramatically improved phase splitting during work-up, which facilitated product isolation and catalyst removal. The optimized biotransformation was successfully demonstrated in a manufacturing trial at 3.5 t scale in an 8000 L reactor. (3R,3aS,6aR)-Hexahydrofuro[2,3-b]furan-3-ol (bisfuran alcohol), a key building block in the synthesis of human immunodeficiency virus (HIV) protease inhibitors such as brecanavir, can be accessed using a number of asymmetric approaches which include lipase resolution.106 At first glance, lipase-catalysed acylation appears to be an attractive possibility for resolution, as there is the potential to remove the undesired alcohol through derivatization whilst leaving the desired enantiomer unchanged for subsequent chemical transformation. However, the desired alcohol is extremely water soluble, which eliminates the possibility of a simple extractive work-up.107 In contrast, highly enantioselective hydrolytic resolution of the racemic acetate, using either PCL or CALB, affords the unwanted enantiomer as an alcohol that can be removed from the desired (R)-acetate on partition between dichloromethane and water.108 During the separation process, a thick emulsion is formed if free enzyme is present. Emulsion formation can be avoided if an immobilized enzyme is used, but enzyme immobilization generally dilutes catalyst activity due to the large quantity of inert support that is required. Thus, high loadings of Novozym 435 (a commercially available form of CALB specifically designed for use in organic solvents) were required to perform the reaction at a reasonable rate, and this led to additional problems such as product absorption and catalyst swelling. By instead


37

employing commercially available ChiroCLEC-PC (a cross-linked crystalline form of PCL), the reaction proceeded rapidly at low loadings (0.05 wt %) comparable to those of the free enzyme, whilst facilitating catalyst recovery and avoiding emulsion formation (Scheme 1.33) (T.C. Lovelace, personal communication).109

O

OAc

H

N

O H

ChiroCLEC-PC O

S

OAc

H

O

+

O H

H

OH

H

O

O N H

N

O S

O

OH

O

(Racemic) O

O

O

O H

H

Brecanavir

Scheme 1.33 Kinetic resolution of a bisfuran intermediate of brecanavir

An example where enzyme-catalysed acylation has been used to good effect was reported by Vaidyanathan et al.110 for the preparation of an androgen receptor antagonist that was being developed as a treatment for alopecia and oily skin. The group were concerned that chromatographic separation of a racemic hydroxynitrile intermediate would afford ultrapure material with an impurity profile that would not be representative of a future commercial process. Enzymatic resolution could provide a practical solution, but enantioselective acylation with commonly used acyl donors like vinyl acetate would afford a neutral product that might be difficult to separate from the starting material and, therefore, also require chromatographic purification. The authors rationalized that, by employing succinic anhydride, previously demonstrated to be an effective acyl donor when used with some lipases,111 an acidic product would result that could then be easily separated from the remaining alcohol by extraction with aqueous base. By screening a variety of lipases in organic solvent for their ability to acylate the racemic hydroxynitrile with succinic anhydride, Novozym 435 was found to yield the best results, affording product in 94–95 % ee at conversions of 47–49 % (Scheme 1.34). After optimization, the reaction was successfully run at 22 kg scale. The immobilized catalyst could be easily isolated by filtration and reused. OH CN (Racemic)

Novozym 435, TBME, 40 – 50 °C O

O

OH

O

CO2H

O

CF3

CN

CN

+ CN

O CN

O

Scheme 1.34 Lipase resolution of a key intermediate in the synthetic route to an androgen receptor antagonist. TBME: tert-butyl methyl ether

Given that resolution can only achieve a maximum yield of 50 %, the approach is inherently inefficient. Additionally, classical resolution and simulated moving bed

38


chromatography can provide attractive alternatives, as development times are frequently shorter, there are no intellectual property or sourcing issues and the techniques are often more accessible to the process chemist. Some examples of where biotransformations have ultimately been replaced by alternative technologies are discussed below. Lotrafiban, a nonpeptidic glycoprotein IIb/IIIa receptor anagonist that was under development as a treatment for the prevention of platelet aggregation and thrombus formation, was initially prepared using an 11-step linear sequence starting from methyl Cbz-Laspartate (Scheme 1.35).112 An overall yield of 9 % and issues with obtaining the product in sufficient enantiopurity led the group to look for an alternative route via the enzymatic NH

CO2tBu HO2C

CO2Me NHCbz F

N

O

HN MeO2C

H2N

N

O

CO2tBu

N

N

HCl. HN

O

O

MeO2C

CO2Me

Lotrafiban

Scheme 1.35 Medicinal chemistry approach to lotrafiban

resolution of a racemic ester intermediate. The ester was screened against a panel of enzymes for hydrolysis activity from which only Novozym 435 efficiently hydrolysed the desired (S)-enantiomer.113 After significant optimization studies using Novozym 435, a process was established where a 100 g L1 slurry of racemic ester in commercial tert-butanol (which is supplied as a mixture containing 12 % water – anhydrous tert-butanol could not be used due to its higher melting point), furnished the desired acid in 43 % yield and >99 % ee (Scheme 1.36). The reaction was performed at 50 °C as a compromise that gave satisfactory substrate concentration

NH HN

O

MeO2C HN MeO2C

N

Novozym 435

N O

N

+

Lotrafiban

O HCl. N HN HO2C

N

HO2C

N O

O

Scheme 1.36 Kinetic resolution of an ester intermediate in the synthetic route to lotrafiban


39

(2.4 g L1) whilst allowing the catalyst to be reused up to 10 times (running at 60 °C, a fivefold reduction in catalyst activity was observed after a single cycle). The undesired enantiomer, remaining as the ester, was separated from the acidic product by selective crystallization and was subsequently racemized and recycled. This route was ultimately run on scale at the site of primary manufacture. In an attempt to find an improved reaction solvent, Roberts et al.113 investigated a number of ionic liquids. Using [BMIM][PF6], an eightfold increase in substrate concentration was observed compared with 88 % v/v tert-butanol, which resulted in a threefold increase in reaction rate and allowed the isolation of acid in comparable yield and enantiopurity to that obtained using the developed process. Ultimately, an alternative route based on asymmetric hydrogenation using a rhodium catalyst employing the Josiphos ligand was identified, but only demonstrated on a 10 g scale before the project was terminated (Scheme 1.37).114

HN

N

Rh(COD)2BF4/L*

O

MeO2C

O

MeO2C L* =

Scheme 1.37 lotrafiban

N

HN

P(iBu)2 Fe P(iPr)2

Asymmetric synthesis of an ester intermediate in the synthetic route to

Lamivudine (also known as Epivir and 3TC) is a potent antiviral drug used in the treatment of HIV and hepatitis B virus (HBV) infections. Although both enantiomers are equipotent antiviral agents, the unnatural enantiomer (with respect to natural nucleosides) is far less cytotoxic, and so a method of selectively accessing the single enantiomer was required. Asymmetric routes to lamivudine have recently been reviewed.115 A number of these are biocatalytic, the most elegant of which is a highly enantioselective kinetic resolution process based on the use of cytidine deaminase from E. coli.116 The process is particularly impressive given that the reaction site is five atoms away from the nearest chiral centre (Scheme 1.38). NH2

O

N O HO

NH2

HN N

O

O

Cytidine deaminase on Eupergit C, pH 7 buffer (166 vols), 32

S (Racemic)

oC,

35 h

HO

N N

O

O

+

N O

HO

S

S Lamivudine

Scheme 1.38

Cytidine-catalysed kinetic resolution of racemic lamivudine

40


Cytidine deaminase was not commercially available, but it is produced by numerous microorganisms and can be induced at high levels in enteric bacteria, such as E. coli, in the presence of cytidine. To overcome the need to add cytidine, mutant strains that express the deaminase constitutively were sought through ultraviolet irradiation of the native microorganism. A selection process was developed to detect strains of interest that took advantage of the fact that both cytidine deaminase and uridine phosphorylase are induced by cytidine as they share the same repressor.117 Thus, any mutant that grows well on uridine in the absence of cytidine is likely to have a defective repressor and express both enzymes constitutively. Using this procedure, the mutant E. coli strain 3732E was developed that gave high deaminase expression independent of cytidine concentration. However, a higher level of expression was required for pilot studies, and so a recombinant strain, overexpressing the deaminase gene from strain 3732E, was developed that produced 80 times more deaminase than the mutant strain. Crude cell extracts of the cytidine deaminase variant, immobilized on Eupergit C, were used on a multikilogram scale. The desired enantiopure product could be selectively extracted by adsorption onto an anionexchange column and isolated in 40 % yield after subsequent recrystallization. The biocatalytic approach was ultimately replaced by the classical resolution of an earlystage intermediate in the final production route. Even so, the deaminase had proven valuable for achieving preclinical supplies.118 Other examples of efficient enzymatic resolutions by reaction at a remote position from stereocentres have been reported, such as the lipase-catalysed resolution of a synthetic intermediate of escitalopram.119 This property of enzymes has also been effectively used to resolve sterically hindered compounds by the introduction of a tether so that the enzyme-catalysed reaction can be performed at an artificially created, but less hindered, remote location. An example is the resolution of tertiary alcohols by the introduction of a glyoxylate ester.120 Most of the examples encountered so far have employed cheap, commercially available enzymes or enzymes that can be readily produced in-house. When a proprietary enzyme, developed by a third party, is used, additional factors such as royalty payments, freedom to operate and single-source supply require consideration. An example is the production of the key (1R,4S)-azabicyclo[2.2.1]hept-5-en-3-one intermediate used in the manufacture of abacavir, another potent reverse transcriptase inhibitor used for the treatment of HIV and HBV infection. Enantiocomplimentary microorganisms (Rhodococcus equi NCIB 40213 and Pseudomonas solanacearum NCIB 40249) were first isolated from the environment under conditions to select for growth on N-acyl compounds as the sole source of carbon and energy.121 Mutant strains of Pseudomonas solanacearum NCIB 40249, hyperexpressing g-lactamase, resulted in a highly enantioselective kinetic resolution process using substrate concentrations of >100 g L1 (Scheme 1.39 where R ¼ H). The process was initially run using whole cells, as the g-lactamase was too unstable to isolate, but this resulted in complex downstream processing. Through further microbial screening, a new lactamase that was sufficiently stable to isolate was identified122 and subsequently cloned (internal presentation from Dow). Using this recombinant lactamase, a highly efficient process was developed that uses 500 g L1 substrate concentrations and a significantly improved workup. In a bid to find a process that employs a commercially available biocatalyst, Mahmoudian et al. rationalized that Boc-protection of the racemic lactam should activate

1.3 Application of Biocatalysis in the Pharmaceutical Industry O

H N

+

41

CO2H

H 2N

γ-Lactamase O

R

HN

N

N

N H2N

R=H R = tBuOC(O)O Savinase (Racemic)

N

N

HO Boc HO2C

O

Abacavir

N

NHBoc

+

Scheme 1.39

Enzymatic kinetic resolution approaches to abacavir

the amide bond towards nucleophilic attack. After screening a variety of commercially available hydrolases towards hydrolysis of this substrate in 1:1 THF/buffer mixtures (to eliminate background hydrolysis), a number of hits were obtained. Of these hits, savinase (protease from Bacillus lentus) proved to be highly enantioselective towards hydrolysis of the undesired enantiomer, leaving the (1R,4S)-Boc-lactam in >99 % ee at 50 % conversion (Scheme 1.39 where R ¼ tBuOC(O)O).123 Savinase and other alkaline proteases are produced in industrial quantities for use in the detergent industry.104b,c Carnell and co-workers have recently applied lipase-catalysed resolution to formally desymmetrize prochiral ketones that would not normally be considered as candidates for enzyme resolution, through enantioselective hydrolysis of the chemically prepared racemic enol acetate.124 For example, an NK-2 antagonist was formally desymmetrized by this approach using Pseudomonas fluorescens lipase (PFL) (Scheme 1.40).125 By recycling the prochiral ketone product, up to 82 % yields of the desired (S)-enol acetate (99 % ee) could be realized.126 This method offers a mild alternative to methodologies such as basecatalysed asymmetric deprotonation, which requires low temperature, and biocatalytic Baeyer–Villiger oxidation, which is difficult to scale up. OAc

O R'N

OAc

O

CN

Ar PFL, n-BuOH CN

Ar

NHR''

+

THF Ar

CN

Ar NK-2 antagonists

O

Ar

CN

Scheme 1.40 Access to NK-2 antagonists by the lipase-catalysed resolution of enol acetates

42


1.3.4.2

Dynamic Kinetic Resolution

As seen in Section 1.3.4.1 (synthesis of lotrafiban), the recycling of an unwanted enantiomer resulting from a kinetic resolution allows theoretical yields of up to 100 % to be achieved, but it can also create a bottleneck in a production process. DKR, where a starting material undergoes racemization in situ, either spontaneously or through the action of a second catalyst, offers a more efficient approach. This technique has been applied, particularly in academia, to the preparation of a broad range of chiral building blocks, and a number of recent reviews are available.127 Odanacatib is currently under clinical development for the treatment of post-menopausal osteoporosis.128 The medicinal chemistry route to the (S)-fluoroleucine moiety, requiring six synthetic steps from an expensive protected aspartic acid derivative and the use of numerous hazardous reagents, was not suitable for scale-up. A more efficient chemoenzymatic approach was instead sought, based on the enzyme-catalysed DKR of racemic 2-phenyl-4-substituted-5(4H)-oxazolones developed by Sih and co-workers.129 The desired racemic azalactone, efficiently produced in a high-yielding, two-pot, four-step process underwent Novozym 435-catalysed ethanolysis in EtOH/TBME in the presence of 20 mol % of triethylamine to furnish ethyl N-Bz-(S)-g-fluoroleucinate in 80 % isolated yield and 95 % ee (Scheme 1.41).130 Unfortunately, benzoyl deprotection of the resultant product could not be effected without significant formation of the desfluoro compound. By instead using the 2-(3-butenyl)-oxazolone, the amino acid derivative was produced in comparable yields, but moderate enantioselectivity (78 % ee). However, deprotection of the 4-pentenamide by hydroxybromination using N,N0 -dibromodimethylhydantoin and trifluoroacetic acid in water/MeCN afforded the desired product in high yield.131 Recrystallization from TBME or isopropyl acetate with H2SO4 afforded the product as the hydrogen sulfate salt in 80 % yield and 97 % ee. This procedure was used to produce >250 kg of API. F

O

OH Base

O N

F

R

O Base

O N

N

R

O

Novozym 435, EtOH O

F

F

R

N H

R

OEt O

F F Deprotect

CF3 N H

OEt

H2SO4. H2N

H N

CN

O

O MeO2S

Odanacatib

Scheme 1.41 Preparation of a g-fluoroleucinate intermediate of odanacatib by enzymecatalysed DKR

In addition to the moderate enantioselectivity, the DKR required one weight equivalent of catalyst to compensate for the background ethanolysis reaction. Furthermore, a significant quantity of hydrolysis product was produced, resulting from the water content of


43

the catalyst that is required for enzyme activity (see Section 1.4). By using a continuous flow format, the biotransformation was greatly improved.132 Not only could the catalyst loading be substantially reduced to 0.05 weight equivalents, but catalyst lifetime was also increased 20-fold due to the absence of shear forces. Product was thus obtained in 90 % yields and 86 % ee in kilogram quantities. The yield of hydrolysis product was reduced, possibly as a result of the catalyst operating at suboptimal water activity due to stripping by solvent. To provide a more efficient route to roxifiban, a drug candidate for the treatment of a range of cardiovascular disorders, Pesti et al. wanted to convert the hydrolytic kinetic resolution of an isoxazoline ester intermediate, using Amano PS30 (PCL), into a DKR.133 Attempts to effect a DKR through adjustment of the reaction pH were unsuccessful even though the ester was prone to base-catalysed racemization via an intramolecular Michael/retro-Michael mechanism. Based on literature precedent for the DKR of -thiophenyl esters, an efficient DKR process was finally established through Amano PS30-catalysed hydrolysis of the n-propyl thioester in triethylamine and aqueous pH 9 buffer solution to furnish the (R)-acid in 80 % yield and >99.9 % ee (Scheme 1.42). OH N

COSPr

O N

COSPr

Trimethylamine

O N

CO2H

Lipase from Ps. cepacia, phosphate buffer CN

CN

CN

H N

AcOH . H2N HN

N

O

NHCO2Bu CO2Me

O

Roxifiban

Scheme 1.42 Enzymatic DKR of a thioester intermediate of odanacatib

Clopidogrel is a potent antithrombotic agent, the chiral portion of which can be accessed from (R)-2-chloromandelic acid. Mandelic acid derivatives are an important class of compound in their own right owing to their use as chiral resolving agents and as building blocks for pharmaceuticals. They can be accessed in enantiomerically pure form by a number of biocatalytic routes, such as nitrile hydrolysis, asymmetric cyanohydrin formation (see Section 1.3.4.5), ketoester reduction (see Scheme 1.53), ester hydrolysis/transesterification,134 O-acetyl hydrolysis135 or hydroxyacid oxidation (Scheme 1.43).136 One of the most attractive biocatalytic options is the nitrilase-catalysed enantioselective hydrolysis of the racemic cyanohydrin. The hydroxyacid is produced directly without need for protection/deprotection steps and cyanohydrins racemize spontaneously at neutral or

44

Biotransformations in Small-molecule Pharmaceutical Development O

OH

R

HO2C

R

R

NC

Hydroxynitrilase

O

HCN

Alcohol dehydrogenase OH

OH

R

HO2C

Monooxygenase

R

NC

Nitrilase

Esterase Esterase

OH

OAc

R

R

HO2C

HO2C OH

R

MeO2C

Scheme 1.43 Some potential biocatalytic approaches to optically pure mandelate derivatives

high pH through the reversible loss of HCN. Another attractive aspect is that, like other hydrolases, nitrilase enzymes require no cofactor. DeSantis et al.137 have reported the discovery of new nitrilases through the screening of genomic libraries created by the extraction of DNA from various environments (metagenomics). In preliminary experiments, using 25 mM mandelonitrile in pH 8 buffer containing 10 % methanol and 0.12 g mL1 of one of these nitrilases, the acid was produced quantitatively with 98 % ee within 10 min. The product was subsequently shown to be (R)-mandelic acid after isolation in 86 % yield. In a parallel reaction, (R)-2-chloromandelic acid was produced at a seventeenth of the rate (Scheme 1.44).

OH

Cl HCN

O

NC

Cl

OH Nitrilase

Cl

HO2C

pH 8

MeO

O

Cl

N S Clopidogrel

Scheme 1.44

Nitrilase-catalysed preparation of a cyanohyrin intermediate to clopidogrel


1.3.4.3

45

Desymmetrization

The initial synthetic route to the antifungal agent posaconazole employed an asymmetric Sharpless–Katsuki epoxidation to afford an (R)-epoxide intermediate in high yield and 88–92 % ee (Scheme 1.45).138 The optical purity could satisfactorily be improved to >98 % ee after one recrystallization of the diol product obtained after ring opening of the epoxide, with retention of stereochemistry, by sodium triazole. Unfortunately, ditosylation and subsequent base-catalysed ring closure of a later triol intermediate gave an almost equimolar mixture of cis- and trans-THF products that required chromatographic separation. OH

OH

F

F

F OH

Sharpless-Katsuki

O

L (+)-tartrate

F

Sodium triazole, DMF

OH N N

F

F

N

CO2Et F

F

O

O O

Na diethyl malonate,

i. MsCl, Et3N, CH2Cl2, 0-5 °C, ii. NaH, DMF

DMF, 50–55 oC

N N

F

N N

F

N

N

OTs

OTs

OH F

F NaBH4, LiCl, EtOH

OH

OH i. TsCl, Et N, DMAP, CH Cl THF 3 2 2–

N N

F

ii. NaH, PhCH3, 100 oC

F O

F

N N

O

+ F

N

N

N N N

40 : 60 Cis/Trans O F

N O

F

N O

N N N N

N N

Posaconazole

Scheme 1.45

HO

Chemical synthesis of posaconazole

This was overcome by acetylation of the same triol intermediate, using Novozym 435 (immobilized CALB) in vinyl acetate and acetonitrile, to afford the monoacetate in 95 % yield and 97 % diastereoselectivity (Scheme 1.46).139 The monoacetate was then readily converted to the desired cis-THF derivative by alcohol activation and cyclization as described above. By performing the desymmetrization on a prochiral diol, a far more efficient asymmetric biocatalytic route was subsequently developed. Enzyme screening found that

46

Biotransformations in Small-molecule Pharmaceutical Development OH

OAc

F

F OH

OH N N

F

CALB

OH

vinyl acetate, MeCN

N N

F

N

Scheme 1.46

OH

N

Lipase-catalysed diastereoselective acetylation of a posaconazole intermediate

CALB was again the favoured catalyst, selectively acetylating the pro-S alcohol (Scheme 1.47). To obtain the desired (S)-monoacetate in sufficient enantiopurity, the reaction was not terminated when all starting material had been consumed, but allowed to run a little further to transform a small portion of monoacetate to diacetate. This resulted in enantioenrichment of the desired (S)-monoacetate by the preferential acetylation of the unwanted (R)-monoacetate to prochiral diacetate. OAc

F F

OAc

OH

OH

F OH

F

CALB

+

vinyl acetate, MeCN

F

I2, NaHCO3, OAc

F

MeCN, 0 °C

O F

I

OAc F

Scheme 1.47

Lipase-catalysed desymmetrization of a posaconazole intermediate

This apparent swap of selectivity is a result of the predictable steric interactions of most commercially available lipases with primary and secondary alcohols and carboxylic acids. In fact, a simple predictive tool, known as the ‘Kazlauskas rules’, has been developed where attack is favoured towards substrates of configuration shown in Figure 1.9.140 These rules are highly predictive for secondary alcohols and less reliable for primary alcohols and carboxylic acids. In the case of the primary alcohols of Scheme 1.47, CALB operates in an antiKazlauskas fashion, resulting in anti-Kazlauskas diol acetylation to produce the (S)monoacetate and anti-Kazlauskas acetylation of the (R)-monoacetate to produce diol (Figure 1.10). In contrast, CALB is observed to act in a Kazlauskas fashion toward the secondary alcohol shown in Scheme 1.34 and the ester shown in Scheme 1.36.

OH M

L

HO CO2H M

L

M

L

Figure 1.9 Kazlauskas rules: preferential action of a lipase on alcohols and carboxylic acids (M and L indicate medium- and large-sized substituents respectively)


HO

HO

F

47

F

AcO

HO

F

F

Figure 1.10 Anti-Kazlauskas action of CALB on the primary alcohol intermediates of posaconazole

The desired S-monoacetate could thus be obtained in 81 % yield and 97 % ee at pilot-plant scale. The tertiary centre could then be constructed by diastereoselective iodocyclization of the resultant monoester, thus removing the need for the Sharpless–Katsuki epoxidation. Diacetate remained unchanged during this step and could be removed at a later stage. Moderate yields of monoacylated product (74–81 %) were initially obtained using vinyl acetate as acylating agent as significant diacetylated by-product formation was necessary to achieve sufficiently high monoacetate enantiopurity. The ultimate route developed for the manufacture of multi-ton quantities of posaconazole used isobutyric anhydride as the acylating agent (Scheme 1.48).141 This more bulky acylating agent proved to be superior, affording >90 % yields of the desired product at low temperature (14 °C) in the presence of NaHCO3 to suppress background reaction and acyl migration respectively. OH OH F

F

OH O

CALB, NaHCO3 O

O

F

O

F

O

Scheme 1.48

Industrial-scale desymmetrization of a posaconazole intermediate

Desymmetrization is not restricted to a single class of enzyme. For example, Madrell et al.142 reported the gram-scale preparation of a key intermediate of the lovastatin lactone through the desymmetrization of 3-(benzyloxy)glutaronitrile using whole cells from Brevibacterium R312. The transformation occurs via a dual nitrile hydratase/amidasecatalysed hydrolysis to afford acid in 65 % yield and 88 % ee (Scheme 1.49). O

HO OBn

OBn Brevibacterium R312

CN

CN

CN

CO2H

O H

O O

Lovastatin

Scheme 1.49 Synthesis of a key hydroxyacid intermediate of lovastatin

48


Using a similar approach, Bergeron et al.143 prepared the side chain of atorvastatin via a nitrilase catalysed desymmetrization of 3-hydroxyglutaronitrile. The dinitrile was prepared in two steps from epichlorohydrin, albeit in moderate yield. A highly enantioselective desymmetrization was then performed using the nitrilase BD9570, developed by Burk and co-workers,144 expressed in a strain of Pseudomonas fluorescens (Scheme 1.50). The enzyme was obtained solely as a soluble, active multimer in excess of 25 g L1 by fermentation, a quantity that represented >50 % of the total cell protein. An advantage of high-level protein expression is greatly simplified downstream processing of the enzyme, a contributing factor to the enzyme cost. In addition, if the enzyme is inexpensive there is no need to recycle, therefore potentially obviating the need for catalyst immobilization. However, reaction workup was problematic due to the high water solubility of the product and the presence of cell debris resulting from the use of crude catalyst.

Cl

O

OH

i. HCN

Cl

CN

OH

NaCN

NC

CN

ii. Base OH

Nitrilase

NC

pH 7.5, 16 h

Scheme 1.50

1.3.4.4

CO2H

H2SO4 EtOH

OH NC

CO2Et

Nitrilase desymmetrization approach to the atorvastatin statin side chain

Asymmetric Ketone Reduction

Microbial reduction has been recognized for decades as a laboratory method of preparing alcohols from ketones with exquisite enantioselectivity. The baker’s yeast system represents one of the better known examples of biocatalysis, taught on many undergraduate chemistry courses. Numerous other microorganisms also produce the ADH enzymes (KREDs) responsible for asymmetric ketone reduction, and so suitable biocatalysts have traditionally been identified by extensive microbial screening. Homann et al.145 have recently reported the identification of a subset of 60 ADH-producing microbial cultures that cut microbial screening time from weeks to days. The advantage of using living microorganisms for bioreduction is that they can be readily sourced from the environment and the cofactors (necessary to regenerate the reduced form of the ADH enzyme and, thus, allowing catalyst turnover) are constantly generated by the intact cellular metabolic machinery. However, reduction using native microorganisms does have several drawbacks. Microorganisms often contain a number of ADHs that can display different or opposite enantioselectivities towards a given substrate. Also, enzymes displaying competing activities might be present or the desired enzyme might not be sufficiently active towards a chosen substrate or poorly expressed by the native organism. Furthermore, most living cells only tolerate low substrate and organic solvent concentrations. For example, Barbieri et al.146 used whole cells from Geotrichum candidum to produce 2 g L1 titres of (S)-chlorohydrin in 90 % yield and 93 % ee. The chlorohydrin can be used as a chiral building block in the synthesis of sertraline, an antidepressant and anorectic agent (Scheme 1.51). To overcome product inhibition, two

1.3 Application of Biocatalysis in the Pharmaceutical Industry OH

O Cl

Cl

G. candidum, XAD-1180 resin

Cl

O NaOH Cl

Cl

Cl

Cl

Cl

O

49

NHMe

O Diethyl malonate Na, dioxane

Cl Cl

Cl Cl Sertraline

Scheme 1.51

Chemoenzymatic approach to sertraline

weight equivalents of the nonionic macroreticular resin Amberlite XAD-1180 was used for in situ product removal. This resulted in a twofold increase in product titre from an unoptimized reaction and both yield and enantioselectivity also increased. Product extraction from large volumes of fermentation broth can be complex, requiring large volumes of organic solvent or solid-phase extraction techniques, which can sometimes greatly reduce or even cancel out the benefits of the biotransformation itself, such as shorter route and environmentally benign conditions. Given the large capital investment required for specialist equipment, the fermentation needs to display considerable production cost benefits over the chemical process to be considered seriously as a route to API manufacture. Partially purified or isolated ADHs offer several advantages: • higher substrate concentrations • higher solvent tolerance • simplified downstream processing. Unlike the whole-cell system, enzymatic reductions require the addition of a hydride donating cofactor to regenerate the reduced form of the enzyme. Depending on the chosen ADH, the cofactor is usually NADH or NADPH, both of which are prohibitively expensive for use in stoichiometric quantities at scale. Given the criticality of cofactor cost, numerous methods of in situ cofactor regeneration, both chemical and biocatalytic, have been investigated. However, only biocatalytic regeneration has so far proven to be sufficiently selective to provide the cofactor total turnover numbers of at least 105 required in production.147 Biocatalytic approaches to cofactor regeneration can be divided into coupled-enzyme methods and coupled-substrate methods.148 In the coupled-enzyme method, the oxidized cofactors (NADþ and NADPþ) are recycled in situ by performing an oxidation reaction using a second enzyme and an inexpensive auxiliary substrate. This second enzyme must employ the same cofactor, but neither enzyme should be able to accept the same substrate.

50


Furthermore, the oxidation reaction needs to be irreversible so as to drive the reduction reaction to completion. NADþ and NADPþ are most frequently recycled using formate dehydrogenase (FDH) and glucose dehydrogenase (GDH) enzymes respectively as the second enzyme. By the introduction of formate and glucose as co-substrates, the oxidized forms of FDH and GDH irreversibly generate carbon dioxide and D-glucono-1,5-lactone respectively, thereby driving the reduction to completion. Alternatively, another ADH can be employed as the second enzyme in the presence of an inexpensive ketone so long as the resultant alcohol can be removed from the reaction mixture in some way as it forms. Davis et al.149 adopted the coupled-enzyme method to access the (S)-hydroxyester (Scheme 1.52) that is subsequently fed into the halohydrin-dehydrogenase-catalysed cyanation process shown in Scheme 1.26. Reaction workup using wild-type enzymes gave an emulsion that settled slowly, thus wasting valuable plant time. Modification of both ADH and GDH enzymes allowed improved separation as well as increased reaction rate and catalyst stability. O

OH

ADH

CO2Et

Cl

CO2Et

Cl

NADP +

NADPH D-glucono-1,5-lactone

D-glucose

GDH

Scheme 1.52

ADH reduction approach to the atorvastatin side chain

Recombinant cells expressing a cloned ADH have also been used in a coupled enzyme method to efficiently produce the (R)-2-chloromandelate intermediate in the synthetic route to clopidogrel in 90 % yield and >99 % ee at 200 gL1 substrate concentration (Scheme 1.53).150 This procedure does not use hydrogen cyanide and, therefore, represents a less hazardous alternative to the nitrilase- and hydroxynitrilase (HnL)-catalysed approaches shown in Scheme 1.44 and Scheme 1.56 respectively. O

OH

Cl

MeO2C

Cl

MeO2C

ADH

NADP+

NADPH D-glucono-1,5-lactone

D-glucose

GDH

Scheme 1.53

ADH approach to the (R)-2-chloromandelate intermediate to clopidogrel

The coupled substrate method is perhaps the simplest approach to asymmetric ketone reduction, using a single recombinant ADH to perform the oxidation of a cheap auxiliary


51

substrate (such as a low molecular weight alcohol) in addition to the desired reduction. By using a large excess of sacrificial alcohol, the reaction can be driven towards formation of the desired reduced product. Montelukast, a leukotriene antagonist used for the treatment of asthma, is produced as a single enantiomer. Asymmetric reduction of the ketone with most hydrogenations and metal hydrides is precluded due to the presence of other sensitive functionality. Using ()-b-chlorodiisopinocamphenylborane (()-DIP-Cl) as the reducing agent at 20 °C, the desired alcohol can be produced in 80 % isolated yield and 99.5 % ee,151 but 1.8 equivalents of this moisture-sensitive and corrosive reagent are required (Scheme 1.54). In light of the need to use stoichiometric quantities of reagent, the development of more efficient catalytic methods has been the subject of extensive research. O Cl

CO2Me

OH

(–)-DIP-Cl (1.8 equivs), THF, Cl

N

CO2Me

N

–20 °C, 4 h

CO2Na

S Cl

HO

N

Montelukast

Scheme 1.54 Preparation of a montelukast intermediate using a chemical asymmetric catalyst

Using a microbial screening strategy, Shafiee et al.152 found that the chiral hydroxyester can be generated from Microbacterium campoquemadoensis in >95 % ee. The whole-cell reaction was optimized to produce 500 mg mL1 product concentrations after 280 h. The ADH responsible was purified and found to be NADPH dependent and active in hexane or DMSO/aqueous mixtures, but no attempt to clone this enzyme has been reported. O Cl

N

Scheme 1.55

CO2Me

OH

ADH, NADPH IPA,toluene, water, 45 oC

Cl

CO2Me

N

Alcohol dehydrogenase preparation of a montelukast intermediate

Ulijn et al. identified an enzyme, capable of enantioselectively reducing the ketone, from their extensive collection of ADH variants; further modification of the hit resulted in a biocatalyst that produces the desired (S)-alcohol in >99.9 % ee at concentrations of 100 gL1 in a solid-to-solid biotransformation,153 where both starting material and product display only sparing solubility in the reaction medium.154 High conversions (>99 %) are achieved by the substrate-coupled method, using 50 % v/v isopropyl alcohol concentrations to drive the reaction by continuous acetone removal (Scheme 1.55). The product can be easily isolated by filtration and washing.

52


Both enantiomers of 1-[3,5-bis(trifluoromethyl)phenyl]ethan-2-ol are of importance in the pharmaceutical industry, and so considerable effort has been expended in their asymmetric synthesis. The (R)-enantiomer is a building block for aprepitant, a neurokinin-1 (NK-1) antagonist used for the treatment of chemotherapy-induced nausea (Figure 1.11).155 Gelo-Pujic et al.156 recently reported the results of a comparison between enzymatic, microbial and chemocatalytic asymmetric reduction of 1-[3,5-bis(trifluoromethyl)phenyl]ethanone. Whereas both biocatalytic methods gave high product ees, both systems only functioned at low substrate concentrations and the enzymatic method gave inferior conversions to the whole-cell system. The chemocatalytic method gave moderate product ees but could be performed at high substrate concentrations and gave high yields. However, the enzymatic approach was only tested using the substrate-coupled method. In sharp contrast, Pollard et al.157 efficiently prepared both alcohol enantiomers with different isolated ADHs using the enzyme-coupled method. For example, using the commercially available ADH from Rhodococcus erythropolis and a GDH cofactor recycling system they produced (R)-alcohol in >98 % yield and >99 % ee at 200 g L1 concentrations on a 25 kg scale. Caution clearly needs to be taken in the proper choice of reaction conditions.

F CF3 H N

O

N

HN N

CF3

CF3

HO

CF3

O

O Aprepitant

Figure 1.11

1.3.4.5

Aprepitant and an (R)-1[3,5-bis(trifluoromethyl)phenyl]ethan-2-ol intermediate

Asymmetrization Using Other Biocatalysts

Another class of biocatalyst of great potential for the preparation of chiral intermediates through asymmetric carbon–carbon bond formation is the HnLs. A range of HnLs are commercially available which are enjoying increasing interest in the pharmaceutical industry. In addition to the nitrilase and ADH approach to the (R)-2-chloromandelate intermediate to clopidogrel discussed earlier (Schemes 1.44 and 1.53), asymmetric cyanation of 2-chlorobenzaldehyde using the crude HnL from Prunus amygdalus (almond meal) has also been reported.158 The reaction is run at low pH (to slow the background reaction), to afford the cyanohydrin in 90 % ee (Scheme 1.56). Several approaches to statin side-chain intermediates have so far been discussed. Whereas these chemoenzymatic approaches provide clear benefits over the chemical processes, they do not harness the true potential of biocatalysis as the biotransformations have simply been inserted into the existing chemical route. Wong and co-workers have developed a more biosynthetic-like approach by using a mutant 2-deoxyribose-5-phosphate aldolase (DERA)

1.3 Application of Biocatalysis in the Pharmaceutical Industry OH

Cl O

Cl

OH

NC

HnL, HCN,

Cl

HO2C

Chemical

Low pH

53

Hydrolysis

MeO

O

Cl

N S Clopidogrel

Scheme 1.56 Preparation of a clopidogrel hydroxyacid intermediates with HnL

(Scheme 1.57).159 Although the natural donor aldehyde is D-2-deoxyribose-5-phosphate, non-phosphorylated donor aldehydes are also tolerated and the enzyme displays some flexibility towards both donor and acceptor. Importantly, as both donor and acceptor substrates are aldehydes, the enzyme can perform sequential aldol reactions allowing the preparation of a key lactol intermediate to the atorvastatin side chain in a single step. Following substantial modification, this approach is now operated on an industrial scale to produce this intermediate in >100 gL1 concentrations.84 OH O

O Cl

+

O

O Cl

Scheme 1.57

O

DERA

+

Cl

O

OH

O O

DERA approach to the atorvastatin side chain

In a recent patent, Hu et al. reported a similar procedure where the acceptor aldehyde contains aminoalkyl substituents in place of chloride.160 Subsequent to lactol oxidation and amine deprotection, these intermediates can directly undergo Paal–Knorr cyclization with the appropriate diketone to produce atorvastatin, thus avoiding the use of cyanide chemistry. The flexibility of DERA enzymes makes them a valuable synthetic tool for the quick access to a range of polyoxgenated products, such as the cytotoxic agent epothilone A (Scheme 1.58).161 Of the known classes of aldolase, DERA (statin side chain) and pyruvate aldolases (sialic acids) have been shown to be of particular value in API production as they use readily accessible substrates.162 Glycine-dependent aldolases are another valuable class that allow access to b-hydroxy amino acid derivatives. In contrast, dihydroxyacetone phosphate (DHAP) aldolases, which also access two stereogenic centres simultaneously,

54

Biotransformations in Small-molecule Pharmaceutical Development OH OH

i. Dowex (H+) OMe

+

O

S

I

O ii. DERA OH

OMe

O

N

S

OAc

HO

N O O

OH HO

O

DERA +

O

PMPO

O

O

Epothilone A OtBu

OH O

Scheme 1.58

OH

OTBSO

DERA approach to the synthesis of epothilone A

have only been of academic interest as they require expensive phosphorylated aldehyde donors and produce phosphorylated products that require subsequent deprotection. This is beginning to change with the discoveries of fructose-6-phosphate aldolase (FSA) that accepts dihydroxyacetone (DHA) as substrate163 and that DHAP aldolase can accept DHA when used in borate buffer due to the transient formation of a borate ester that mimics phosphate.164 As more enzyme kits become commercially available, the screening for a suitable catalyst can now be performed in a matter of hours rather than days or weeks. Furthermore, both the screening and biotransformation can be performed by nonspecialists. This increases the likelihood of uptake of a biocatalytic process, as a proof of concept can be more readily obtained without the commitment of considerable resource. For these reasons, the use of ADHs by pharmaceutical companies has increased considerably in recent years.

1.4

Enzymes in Organic Solvent

Biocatalysis has traditionally been performed in aqueous environments, but this is of limited value for the vast majority of nonpolar reactants used in chemical synthesis. For a long time it was assumed that all organic solvents act as denaturants, primarily based on the flawed extrapolation of data obtained from the exposure of aqueous solutions of enzyme to a few water-miscible solvents, such as alcohols and acetone, to that of all organic solvents.165 This assumption has since been swept aside and it is now recognized that a broad range of enzymes retain their activity on exposure to organic solvents or organic solvent–water mixtures. The addition of organic solvent allows the coupling of the exquisite selectivities observed from traditional approaches with numerous other advantages, such as: • • • •

increased concentrations of nonpolar reactants; enablement of reactions that have unfavourable thermodynamic equilibria in water; enhanced biocatalyst stability towards heat and autolysis; compartmentalization of substrate/product from enzyme (reduced substrate/product inhibition); • modification of enzyme selectivity; • selective inhibition of competing enzymes;

1.4 Enzymes in Organic Solvent

• • • •

55

reduced background reaction; improved workup; better integration into synthetic routes; greater potential for tandem chemoenzymatic processes.

The field of biocatalysis in organic media is now of considerable industrial importance, enjoying widespread application, particularly in the preparation of enantiopure intermediates.166 Enzyme catalysis in nonconventional media can be divided into a number of different categories depending on whether the aqueous and organic phases are miscible or immiscible and whether the biocatalyst is dissolved or not. In this section, only ‘free’ enzymes will be considered. Thus, the field can be simplified to just two categories, depending on whether the solvent is water miscible or immiscible (systems employing water-immiscible solvents, where water is present in quantities that are below its solubility limit, have been considered as monophasic): 1. monophasic biocatalysis 2. biphasic biocatalysis. The state of the catalyst (homogeneous or heterogeneous) is dictated by the relative quantities of solvent and water used. 1.4.1

Monophasic Biocatalysis

The structural integrity of enzymes in aqueous solution is often compromised by the addition of small quantities of water-miscible organic solvents.167 However, there are numerous examples, particularly using extremophiles,168 where enzymes have been successfully employed in organic solvent–aqueous mixtures.166b A good example is the savinase-catalysed kinetic resolution of an activated racemic lactam precursor to abacavir in 1:1 THF/water (Scheme 1.39). The organic solvent is beneficial as it retards the rate of the unselective background hydrolysis. The use of water-miscible organic solvent–water mixtures is a particularly attractive method for use with cofactor-dependent enzymes due to its simplicity. The high water content can allow dissolution of both enzyme and cofactor, whilst the water-miscible solvent can provide a dual role in both substrate dissolution and as a cosubstrate for cofactor recycling (substrate-coupled cofactor recycling).148 The asymmetric reduction of a ketone intermediate of montelukast using an engineered ADH in the presence of 50 % v/v isopropanol offers a powerful demonstration of this methodology (Scheme 1.55). It might be expected that in miscible organic solvent–water mixtures of increasing organic solvent content, the structural integrity of many enzymes will progressively diminish due to loss of essential hydrogen bonding. In fact, this is not the case, as demonstrated by Griebenow and Klibanov,165 who used Fourier-transform infrared spectroscopy to assess the effect of acetonitrile–water mixtures (0–100 %) on the secondary structure of lysozyme. Rather than a gradual loss in secondary structure with increasing organic solvent content, they observed an inverse bell-shaped relationship, with maximum -helicity occurring at both high water and high organic solvent content. Reduced enzyme solubility at high organic solvent content might have provided an attractive rationale, but this was not supported by the data. A similar trend was observed using Bacillus subtilisin

56


protease (also known as subtilisin Carlsberg) and other water-miscible organic solvents. The authors concluded that enzyme denaturation increases as the organic solvent content increases. At the same time, a decline in water content reduces conformational mobility so that the enzyme becomes kinetically trapped in an active conformation. In addition to the retention of structural integrity in neat organic solvents, Klibanov and co-workers demonstrated that a diverse range of enzymes, from hydrolases and peroxidases to cofactor-dependent alcohol oxidases and ADHs, also retain activity.67 This pioneering work single-handedly led to the popularization of biocatalysis in neat organic solvent. Recent literature has shown that nonaqueous biocatalysis is not limited to traditional organic solvents, with examples that employ ionic liquids169 and supercritical fluids170 now widespread. Reaction in organic solvent has also led to the discovery that some enzymes display promiscuity towards reaction type as well as substrate type,171 with the HnL-catalysed asymmetric Henry reaction,172 and the lipase-catalysed Michael-type addition of thiols to ,b-unsaturated enones providing some recent examples.173 Enhanced rigidity of enzymes in nonaqueous media also imparts greater thermostability, allowing reactions to be run at temperatures of up to 100 °C over prolonged time periods.174 For example, the kinetic resolution of a key intermediate in the synthesis of lotrafiban using Novozym 435 as catalyst (Scheme 1.36) can be performed at temperatures of 70 °C over prolonged reaction times without enzyme degradation. However, a lower temperature of 50 °C was employed in the final production route due to limitations of the immobilization technique used rather than the enzyme. In the 88 % tert-butanol–12 % water solvent mixture, required to provide sufficient substrate solubility, substantial enzyme desorption from the support at higher temperatures limited reuse of this expensive catalyst. Efficient biocatalysis in neat organic solvent depends on the careful choice of the method of ‘dehydrated’ enzyme preparation and solvent used. Optimization of these factors towards a given transformation is often known as ‘catalyst formulation’ and ‘solvent, or medium, engineering’ respectively, both of which will be briefly discussed below. ‘Catalyst engineering’ which also provides a powerful method of improving activity and stability, is discussed in Chapter 2. 1.4.1.1

Catalyst Formulation

A requirement of biocatalysis in neat organic solvent is the use of a dehydrated form of an enzyme that displays the desired activity. A number of techniques are available for the preparation of dehydrated enzymes, some of which are discussed in a recent review by Griebenow and Barletta.175 The techniques that have been most commonly used are: • lyophilization • precipitation • immobilization (see Section 1.5). The resultant dehydrated enzyme preparations often display comparable activity to untreated enzyme when reconstituted in aqueous buffer. However, in the case of many enzymes, activity in a suitable neat organic solvent can be three to five orders of magnitude lower than in water. This was recognized by Klibanov early on in the


57

development of the field, and so many of the basic principles leading to reduced efficiency have been elucidated. These have been extensively reviewed and will only be briefly discussed here.176,177 A major cause of suboptimal activity in organic solvent results from the removal of ‘essential water’ during enzyme dehydration. All enzymes require some water in order to retain activity through the provision of conformational flexibility.178 Particularly in the case of lipases, the amount of water can be so low that it appears that none is required. For example, following the development of suitable techniques to analyse low water concentrations,179 it has been reported that the lipase from Rhizomucor miehei retains 30 % of its optimum activity with as little as two or three water molecules per molecule of enzyme.180,181 Owing to the apparent absence of water in some exceptional cases, the term ‘biocatalysis in anhydrous solvent’ is commonly used, although in the vast majority of cases a monolayer of water is required for optimal activity (although this is often still well below its solubility limit in water-immiscible solvent).67 Numerous ‘tricks’ have been developed to retain activity of the dehydrated enzyme preparation. Activity can be dramatically enhanced by adding a small quantity of water to the enzyme prior to use,182 but this can be detrimental in transformations where it can participate as a reactant, particularly where the reagents are expensive. Retention of activity without the need to partially rehydrate has, therefore, been the focus of intensive investigation. Some effective strategies, such as co-lyophilization in the presence of lyoprotectants (sugars or hydrophilic polymers) and the use of additives such as crown ethers, substrate or transition-state analogues (molecular imprinting) or inorganic salts, have recently been reviewed by Serdakowski and Dordick.177 Some of these techniques can lead to dramatic changes in enantioselectivity and activity.183 The ionization state of polar (ionogenic) residues of the dehydrated enzyme preparation can also have a substantial impact on conformation and, hence, on activity in organic solvent. The ionization state can be optimized through pH control of the aqueous solution from which the enzyme was last in contact. Commonly referred to as the ‘pH memory’ effect, optimum activity in organic solvent is usually attained by preparing the dehydrated enzyme from an aqueous solution of optimal pH for enzyme activity in conventional media. In many cases, charged species are generated during the course of a transformation that can affect the enzyme ionization state. This can be controlled through the addition of solid-state buffers to the reaction mixture.184 Because enzymes are insoluble in organic solvent, mass-transfer limitations apply as with any heterogeneous catalyst. Water-soluble enzymes (which represent the majority of enzymes currently used in biocatalysis) have hydrophilic surfaces and so tend to form aggregates or stick to reaction vessel walls rather than form the fine dispersions that are required for optimum efficiency. This can be overcome by enzyme immobilization, as discussed in Section 1.5. 1.4.1.2

Solvent Engineering

Enzyme activity varies greatly depending on solvent choice, as illustrated by Zaks and Klibanov185 for the transesterification of tributyrin and heptanol by three different lipases. Using these data, Laane et al.186 found that enzyme activity correlates closely with solvent hydrophobicity (log P) for the lipases from Mucor sp. (MML) and Candida cylindracea

58


Figure 1.12 Transesterification activity of PPL, CCL and MML in various organic solvents

(CCL – now known as lipase from Candida rugosa (CRL)) but not porcine pancreatic lipase (PPL) (Figure 1.12). It was postulated that the differences in enzyme activity observed primarily result from interactions between enzyme-bound water and solvent, rather than enzyme and solvent. As enzyme-associated water is noncovalently attached, with some molecules more tightly bound than others, enzyme hydration is a dynamic process for which there will be competition between enzyme and solvent. Solvents of greater hydrophilicity will strip more water from the enzyme, decreasing enzyme mobility and ultimately resulting in reversible enzyme deactivation. Each enzyme, having a unique sequence (and in some cases covalently or noncovalently attached cofactors and/or carbohydrates), will also have different affinities for water, so that in the case of PPL the enzyme is sufficiently hydrophilic to retain water in all but the most hydrophilic solvents. The impact of water on enzyme activity is powerfully demonstrated by the chymotrypsincatalysed transesterification of ethyl N-acetyl-L-phenylalaninate with propanol. In dry acetone, the reaction is over 7000 times slower than in dry octane. However, by adding 1.5 % v/v water to acetone, the reaction rate dramatically increases to two-thirds the rate of that in dry octane.67a Zaks and Klibanov also demonstrated the effect of water stripping on enzyme activity by incubating chymotrypsin in various organic solvents and then assessing the resulting enzyme water content. Activity in the different organic solvents was found to correlate well with water retained by the enzyme. Halling was able to rationalize such findings by realizing that a given enzyme requires a defined quantity of water to attain optimal activity. This can be expressed in terms of thermodynamic water activity, which essentially describes the amount of water bound to the enzyme.187 Thus, optimum chymotrypsin activity in acetone is realized at the same thermodynamic water activity as that in


59

octane even though the total water content of each system is very different. However, at comparable water activity, variations in optimum enzyme activity observed in each solvent show that the direct effect of solvent on the enzyme is also an important factor which may account for the activity deviations from the activity/log P relationship seen in Figure 1.12. The choice of organic solvent can also have a dramatic effect on selectivity.166a In contrast to enzyme activity, in the majority of examples reported there appears to be no correlation between solvent physical properties and enantioselectivity. In fact, investigating the effect of various solvents towards a number of lipases, Secundo et al.188 also found that the optimal solvent differed with both enzyme and substrate. A number of theories have been postulated in order to explain these effects in individual cases, but none has any general predictive value.183b This is somewhat intriguing given that differences in enantioselectivity simply relate to a change in the relative rate of conversion of each enantiomer. Reaction in organic solvent can sometimes provide superior selectivity to that observed in aqueous solution. For example, Keeling et al.189 recently produced enantioenriched -trifluoromethyl--tosyloxymethyl epoxide, a key intermediate in the synthetic route to a series of nonsteroidal glucocorticoid receptor agonist drug candidates, through the enantioselective acylation of a prochiral triol using the lipase from Burkholderia cepacia in vinyl butyrate and TBME (Scheme 1.59). In contrast, attempts to access the opposite enantiomer by desymmetrization of the 1,3-diester by lipase-catalysed hydrolysis resulted in rapid hydrolysis to triol under a variety of conditions. F3C HO

OH OH

Lipase, vinyl butyrate

F3C HO

OH O

Pr

O

CF3 O

Pr

TBME 85% yield 92% ee

R2 O

R1 OH N F3C

Scheme 1.59

1.4.2

N N

H N O

O

O

R

NH2

Synthesis of nonsteroidal GR agonists

Biphasic Biocatalysis

Biocatalysis in biphasic mixtures of water-immiscible organic solvent and water involves the transfer of low concentrations of substrate from the organic to aqueous phase during agitation. The substrate then undergoes transformation before returning to the organic phase. The partition of substrate/product between the two phases is independent of their ratio and so the volume of the organic phase can be much greater than the aqueous phase, allowing high-intensity transformations to be achieved whilst simultaneously minimizing exposure of the enzyme to organic species. The technique is particularly valuable for transformations in which the enzyme is sensitive to inhibition by high concentrations of substrate or product and transformations where cofactor recycling is required. Biphasic conditions can also be used to suppress background reaction. HnL-catalysed asymmetric addition of cyanide to aldehydes and ketones provides an important example,

60


allowing chiral intermediates to APIs such as clopidogrel to be accessed in excellent enantiopurity (Scheme 1.56). However, whereas the biphasic method of controlling background reaction works well with nonpolar substrates, it is less effective with polar, water-soluble substrates such as 3-pyridinecarboxaldehyde. Such substrates require transformation under nearly anhydrous conditions where, unfortunately, HnLs rapidly deactivate. Faced with this issue, Roberge et al.190 have recently reported that HnLs, immobilized as cross-linked enzyme aggregates (CLEAs), retain their activity in nearly anhydrous conditions (see Section 1.5.2 for further details of CLEAs). Using two different commercially available HnL CLEAs they were able to produce either of the enantiomers of 3-pyridinecarboxaldehyde cyanohydrin in moderate to high yield and >90 % ee in dichloromethane containing just 0.18 % water. The solvent present in biphasic reactions can still have an effect on the enzyme even though the enzyme functions primarily in an aqueous microenvironment. A particularly dramatic example is the lipase AH (lipase from Burkholderia cepacia)-catalysed desymmetrization of prochiral 1,4-dihydropyridine dicarboxylic esters, where either enantiomer can be accessed in high enantioselectivity by using either water-saturated cyclohexane or diisopropyl ether (DIPE) respectively (Scheme 1.60).191 The acyl group used in acylation and deacylation can also have a dramatic effect on enantioselectivity.134 NO2 O lipase AH, cyclohexane, water

NO2 O

O O

O

O

HO

O

O

N H R-ent 87% yield, 89% ee

O

O

O

O

O NO2

N H lipase AH, DIPE, water

O

O

O O

O

OH N H

S-ent 88% yield, >99% ee

Scheme 1.60 Resolution of a prochiral 1,4-dihydropyridine dicarboxylic ester with lipase AH in the presence of cyclohexane or DIPE

In conclusion, by using organic solvents, biotransformations can achieve productivities suitable for pharmaceutical manufacture. Biocatalysis under organic solvent–aqueous conditions can be applied to a broad range of enzymes as the methodology is compatible with cofactor recycling, whereas biocatalysis in nearly anhydrous solvent facilitates numerous transformations that are thermodynamically disfavoured in the presence of water, although limited to use with enzymes that do not require cofactors, particularly hydrolases. In selecting an appropriate solvent, it is necessary to screen each new biotransformation on a case-by-case basis to ensure that optimum enzyme activity, stability and selectivity are

1.5 Enzyme Immobilization

61

achieved. For optimal activity under nearly anhydrous conditions, attention should also be paid to water activity and the dehydrated enzyme formulation used. Water stripping is particularly important to consider when setting up a continuous process.

1.5

Enzyme Immobilization

Ballesteros et al.192 defined immobilized biocatalysts as ‘enzymes, cells or organelles (or combinations of these) which are in a state that permits their reuse’. Enzyme immobilization represents only a small part of this field, but is the most commonly employed in pharmaceutical production. Immobilized enzymes are frequently used in biocatalysis to overcome limitations such as: • • • • • • •

insufficient stability towards temperature, pH, shear stress or autolysis; necessity to recycle the enzyme for economical reasons; biological contamination of the product causing complex downstream processing; emulsion formation during product extraction; poor catalyst dispersion in the reaction mixture; insufficient activity; inappropriate form if required for a continuous process. Where immobilization is necessary, any resulting biocatalyst should be:

• • • • • • • • •

toxicologically safe; low cost; sufficiently active and selective; chemically and thermally stable under process and storage conditions; insoluble towards the reaction solvent; mechanically strong; of uniform particle size; resistant to microbial attack; reusable.

Numerous different immobilization methods have been reported that take advantage of various enzyme properties such as size, chemically reactive functionality, ionic groups or hydrophobic domains.193 Based on these properties, enzyme immobilization can be split into three main classes (which are also applicable to the immobilization of cell cultures): • noncovalent attachment; • covalent attachment and cross-linking; • entrapment. In spite of the immense quantity of available literature, it can still be a challenge to determine which immobilization technique is suitable for a particular application, and so it is usually necessary to test a number of options on a case-by-case basis. 1.5.1

Noncovalent Attachment

Noncovalent attachment is a popular method of immobilization, and numerous different support materials have been employed, ranging from organic supports, like cellulose,

62


chitin, ion-exchange resins and polyacrylamide, to inorganic supports, such as celite, salts, zeolites or even iron particles. However, the technique is disfavoured for industrial applications as the enzyme is weakly bound and, therefore, prone to leaching, potentially leading to product contamination and inefficient recycling. Many lipases are commercially available in a noncovalently immobilized form either adsorbed onto celite, which aids dispersion in organic solvent, or onto a hydrophobic support such as accurel. As a result, noncovalently immobilized lipases are frequently employed, in spite of the above limitations, owing to their availability. Lipase immobilization on hydrophobic supports is particularly useful, as it takes advantage of the unique property of this enzyme class towards interfacial activation at the surface of oil droplets.194 Unlike other enzymes, most lipases contain what is often referred to as a lid or flap that masks the active site. This lid is hydrophilic on the external surface and hydrophobic on the internal surface, so that in aqueous solution the lipase exists in an equilibrium lying primarily towards the inactive or closed form. On adsorption to an oil droplet, the flap undergoes a conformational change to the ‘open form,’ resulting in activation. As discussed in Section 1.4, enzymes are more rigid in organic solvent and so the lipase can be trapped in the form that was predominant in the aqueous solution from which it was last in contact.195 On immobilization, the hydrophobic support itself can mimic an oil droplet, resulting in hyperactivation of the lipase. It is not uncommon for an immobilized lipase to display greatly enhanced activity over that of the free enzyme. 1.5.2

Covalent Attachment and Cross-linking

Immobilization of an enzyme through covalent attachment is a widely used technique, as the catalyst can be used in either aqueous or organic media without leaching and provides a suitable catalyst form for use in multipurpose apparatus or more specialized equipment such as a continuous reactor. Covalent attachment is usually achieved via attack from nucleophilic groups of the enzyme onto electrophilic moieties on the support (although the reverse has also been reported). Given that most enzymes have numerous reactive substituents (Table 1.2), multipoint attachment to the support can occur, which can have a significant stabilizing effect. A drawback of this technique can result from the formation of covalent linkages in or near to the enzyme active site, causing deactivation. However, this outcome can usually be circumvented by using another of the many alternative supports available. Table 1.2

Reactive functionality of amino acid residues frequently present in proteins

Functional group

Amino acid

Primary amine Thiol Carboxylic acid Phenol Guanidine Imidazole Disulfide Indole Thioether Alcohol

L-Lysine

and N-terminus

L-Cysteine L-Aspartate, L-glutamate L-Tyrosine L-Argenine L-Histidine L-Cystine L-Tryptophan L-Methionine L-Serine, L-threonine

and C-terminus

1.5 Enzyme Immobilization

63

Eupergit C and, more recently, Sepabeads EC-EP are mesoporous supports that have proven to be of particular importance in pharmaceutical production. Both are highly hydrophilic macroporous resins, containing high densities of epoxide groups on the surface. Available as beads of 100–250 mm in diameter and 20–40 nm pore diameter, these resins display high chemical and mechanical stability, tolerating a wide range of pH and solvents. About 60 mg of purified enzyme per gram of resin can generally be immobilized onto Sepabeads EC-EP, under extremely mild conditions, using enzyme dissolved in buffers of high salt concentration. An initial rapid adsorption takes place followed by slower covalent bond formation, after which the remaining epoxides (as much as 99 % of the original groups) can be opened or ‘capped’ using a nucleophilic species. Crude enzyme preparations can also be used, as other cell debris will either irreversibly bind to the support along with the enzyme or can be easily washed away after immobilization is complete. To exemplify the mildness and robustness of this technique, 85–90 % of PGA active sites have been reported to remain competent following immobilization to Eupergit C.196 Furthermore, the immobilized catalyst lost only 40 % of its activity over >800 cycles. Covalent enzyme attachment to an inert support is inherently inefficient, as enzyme activity is diluted and additional material costs are incurred. An attractive alternative that circumvents both of these issues is to cross-link enzyme molecules together using a bifunctional linker. This technique gained huge popularity with the emergence of crosslinked enzyme crystals (CLECs).197 CLECs are produced by crystallization of purified enzyme and subsequent cross-linking, usually with glutaraldehyde, which is an FDAapproved fixing agent for the immobilization of glucose isomerase used in high-fructose corn syrup production.198 CLECs proved to be excellent biocatalysts, displaying high activity, stability and separation properties, as demonstrated by their use in the resolution of the bisfuran alcohol intermediate of brecanavir (Scheme 1.33). Unfortunately, enzyme purification and crystallization can be labour intensive to develop and inefficient, resulting in an extremely active but highly expensive catalyst. This led to poor uptake of the technology and withdrawal of CLECs from the marketplace. More recently, CLEAs have been introduced. They provide many of the positive attributes of CLECs but can be rapidly prepared from partially purified enzyme preparations with minimal technical expertise.199 Essentially, their preparation involves enzyme precipitation (see Section 1.4.1.1) with in situ cross-linking, or vice versa. Glutaraldehyde is usually employed as the cross-linking agent, although bulkier linkers, such as dextran polyaldehyde, have been successfully used where cross-linking with the smaller reagent results in activity loss through interaction with the enzyme active site.200 1.5.3

Entrapment

Entrapment involves the physical confinement of an enzyme in a semipermeable matrix, in much the same manner as nature handles soluble enzymes.201 This should represent an extremely mild method of immobilization, as the enzyme remains free, albeit confined to a small space. Two techniques, which at first sight appear unrelated, have been well utilized: • entrapment in a polymer matrix; • entrapment behind a membrane.

64


Entrapment in polymeric matrices is a variation of noncovalent attachment where the support is instead generated in the presence of the enzyme. A particularly popular entrapment technique is sol–gel encapsulation, where the enzyme is trapped within an SiO2 matrix formed by acid- or base-catalysed hydrolysis of tetraalkoxysilanes in the presence of enzyme.202 The technique can be tuned to provide the appropriate microenvironment for each enzyme in much the same way as can be done with other immobilization methods.203 Pharmaceutical production generally uses multipurpose equipment, and so entrapment behind a membrane would require significant capital expenditure on specialized equipment. In spite of this, the use of membrane reactors in biocatalysis represents an efficient method of enzyme immobilization, given the large molecular weight difference between enzymes (10–150 kDa) and most substrates (300–500 Da). The reader is referred to some recent reviews on the topic.204 In summary, enzyme immobilization is extremely important in the scale-up of many biocatalytic processes. The preferred method for pharmaceutical production involves covalent binding through cross-linking or attachment to a support. Noncovalent attachment is less attractive, but it is heavily utilized owing to the commercial availability of industrial quantities of some enzymes immobilized using this technique.

1.6

Green Chemistry

The use of biocatalysis in the manufacture of APIs can address some of the 12 principles of green chemistry set out by Anastas and Warner.205 For example, biocatalytic processes can: • • • • •

increase atom efficiency; operate under mild conditions; reduce protection/deprotection steps; avoid the use of stoichiometric reagents; avoid the use of toxic/hazardous chemistry.

However, these statements are generalizations, and it is not necessarily true to say that all biotransformations will be greener than the chemical alternative. Therefore, it is important to analyse each comparison objectively on a case-by-case basis using a multivariate process to take into account the complexity of the analysis. Designing greener processes involves, for example: • designing efficient processes that minimize the resources (mass and energy) needed to produce the desired product; • considering the environmental, health and safety profile of the materials used in the process; • considering the environmental life cycle of the process; • considering the economic viability of the process; • considering the waste generated in the process, both in nature and quantity, whether it is hazardous, benign, can be recycled or recovered and used in this or another process. It is not easy or straightforward to determine how green a process is, and there have been a number of different approaches taken. Sheldon’s E-factor was one of the first measures of greenness proposed in the 1980s, to highlight the amount of waste generated in order to

1.6 Green Chemistry

65

produce 1 kg of chemical product across different branches of the chemical industry.206 Simply put, the higher the E number, the more waste is generated to produce 1 kg of product. Within the pharmaceutical industry there have been other variations of measuring the mass efficiency, such as the mass intensity proposed by Constable et al.207 and the process mass intensity proposed by the ACS GCI Pharmaceutical Roundtable.208 Measuring greenness is not just about determining the quantity of waste; one must also consider the efficiency of the chemistry or biochemistry (atom efficiency, reaction mass efficiency) and the nature of the materials involved as reagents, solvents and as waste.209 One should also consider the process conditions used, all within the context of the 12 principles of green chemistry. The next factor to take into account when trying to evaluate the greenness is the environmental life cycle impact of the materials used in the process. Determining the life cycle for every material used in a pharmaceutical synthetic process is a complex task, as often the life cycle data for every material is just not available. However, GSK have developed a methodology and a tool to enable good estimations of the life cycle impacts so that comparisons between different development options can be made.210 Data have recently been added to the tool to enable life cycle comparisons for routes using enzymes as catalysts or involving a fermentation step. GSK have also developed a framework for analysing and comparing two processes based upon the suite of metrics discussed above.211 This framework was used as the basis for a comparison of the environmental, health, safety and life cycle (EHS and LCA) impacts of the chemical (Scheme 1.11) and two enzyme biocatalytic (Scheme 1.12) 7-ACA processes, recently reported by Henderson et al.55 The measures used accounted for the chemical and process efficiencies, the nature of the materials used and waste generated, as well as determining the overall life cycle environmental impacts from ‘cradle to gate’ of each process. This analysis showed that the bioprocess could be classed as ‘greener’ when compared with the purely chemical process. The chemical process uses more hazardous materials and solvents, and requires about 25 % more process energy than the enzymatic process. When accounting for the cradle-to-gate environmental life cycle, the chemical process has a larger overall environmental impact, mainly derived from the production of raw materials. In comparison with the enzymecatalysed process, the chemical process uses approximately 60 % more energy, about 16 % more mass (excluding water), has double the greenhouse gas impact and about 30 % higher photochemical ozone creation potential and acidification impact. Only the yield of the chemical process was higher, showing that yield is not a good measure of greenness, which reinforces the message that it is important to take a more holistic view, since assessing greenness is a multivariate and complex process. One of the aims of the analysis was to develop a methodology and framework for objective comparisons of two very different types of synthetic process, which could then be applied to other different systems. A secondary aim was to test the hypothesis that biotransformations are greener than chemical transformations. By the application of such rigorous and academic analyses one can test this hypothesis for a number of different systems, including once-through fermentations and enzyme-catalysed systems, where the amounts of waste generated will be significantly different. To celebrate the fifteenth anniversary of his E-factor, Sheldon compared different measures of greenness212 with the E-factor and reminds us of the value of the headline number, which challenges those in the pharmaceutical industry to improve the efficiency of

66


pharmaceutical processes by moving away from continually using stoichiometric reagents towards catalytic reagents. While it is true to say that the absolute volumes of waste are low compared with fine chemicals or petrochemicals, the challenge remains valid today that the pharmaceutical industry has the opportunity to embrace catalytic technology as one way to improve the mass efficiency of processes. The application of biocatalytic technology in the pharmaceutical industry is one way of addressing that challenge.

1.7

Future Perspectives

Biocatalysis contributes significantly to the generation of APIs through the supply of chiral building blocks from the fine chemical industry. In contrast, there is a clear underutilization within the pharmaceutical industry, where it could provide more efficient and greener methods of late-stage intermediate and API production. The ACS GCI Pharmaceutical Roundtable recently set out to prioritize the areas of chemical synthesis where improved methodology would realize the greatest beneficial impact on pharmaceutical production. This resulted in the publication of a ‘wish list’ of currently utilized transformations that require better reagents and aspirational transformations that would provide shorter routes were they available (Table 1.3).16 A recent categorization of biotransformations by Pollard and Woodley12 (Figure 1.13), based on the availability of commercial enzymes, together with the examples given in this book demonstrate that biocatalysis can meet many of these pharmaceutical needs as shown by the highlighted entries in Table 1.3. Table 1.3 List of key areas of green chemistry of importance to the pharmaceutical industry (in ascending order); areas where biocatalytic precedent exists are given in bold. Reactions currently used but better reagents preferred

More aspirational reactions

Amide formation avoiding poor atom economy reagents

CH activation of aromatics (cross coupling reactions avoiding the preparation of haloaromatics) Aldehyde or ketone þ NH3 þ ‘X’ to give chiral amine Asymmetric hydrogenation of unfunctionalized olefins/enamines/imines New greener fluorination methods

OH activation for nucleophilic substitution Reduction of amides without hydride reagents

Oxidation/epoxidation methods without the use of chlorinated solvents Safer and more environmentally friendly Mitsunobu reactions Friedel–Crafts reaction on unactivated systems Nitrations

N-Centred chemistry avoiding azides, hydrazine etc. Asymmetric hydramination Green sources of electophilic nitrogen (not TsN3, nitroso, or diimide) Asymmetric hydrocyanation

1.7 Future Perspectives Established Chemistries XH

O

CN

R″

X R

R′ R″ R OH R, R′, R″ = Alkyl or aryl X = O, N or S O

Expanding Chemistries

O

Lipase/Protease

+

CO2H

Nitrilase

R'

R

R′

R′

R

R, R′, R″ = Alkyl or aryl ADH

OH

O

R′

R

R′

OH CN R′ R

Hydroxynitrilase

+ R

67

HCN

R′

R

Emerging Chemistries O

NH2

Transaminase

R′ R R = Alkyl or aryl R′ = Alkyl, aryl or CO2R

R

R′

R

R′

O

Monooxygenase

R′

R

O

O

O Monooxygenase

Epoxide hydrolase

O R′

R

OH

R

OH

O

R″

N-oxidase

O

R

R′′′

OH O

O Aldolase

+ R′

R″

R′

OH R

R′

R″

R

R′′′ Monooxygenase

R″

R′

R′′ O

N

R

R′

R′′

R′

H N

R

Enoate reductase

R

R

R′

R

O R

O +

OH

Decarboxylase R′

R′

R O

Figure 1.13 Status of various biotransformations (not exhaustive). (Reprinted from Pollard, D.J. and Woodley, J.M. Biocatalysis for pharmaceutical intermediates: the future is now. Trends Biotechnol. 2007, 25, 66–73 with permission from Elsevier.)

Routes to APIs are predominantly designed by synthetic organic chemists who are well versed in the adoption of new technologies. To maximize uptake of biocatalytic techniques, the most efficient approach is to provide them with reasonably priced kits of enzymes that can easily be used without specialist knowledge. Greater availability of comprehensive commercial kits with diverse applications and better tools to predict improved biocatalyst properties in silico should diminish the current perception by many chemists that enzymes are exotic catalysts only to be used as a last resort. However, this expansion requires significant investment from specialist enzyme producers, many of whom subsequently base their business models on the generation of royalties from the use of their proprietary biocatalysts or biocatalytic processes. The use of proprietary enzymes in pharmaceutical production can be cost effective where a biocatalyst is involved in an asymmetric or regioselective transformation if traditional chemical approaches generate substantial waste or require additional steps, but is probably precluded for achiral transformations such as the replacement of an atom-inefficient coupling reagent for amide bond

68


formation. However, as the field matures and these enzymes become cheaper, such applications should become competitive. The above applications consider biocatalysis from the perspective of the synthetic organic chemist rather than the biochemist. Slotting single-step biotransformations into chemical syntheses is unlikely to use biocatalysis to its full potential. Undoubtedly, isolated enzymes offer an attractive solution to rapid biocatalyst identification, and advances in molecular biology and biotransformation technology have provided a number of techniques by which hits can be modified to fit a required process, or vice versa. However, there is also a significant cost associated with the isolation of enzymes at scale. It is far more attractive to use crude lysates or whole cells; but, as shown in previous sections, these have their own disadvantages. The use of crude lysates can increase downstream processing complexity, and alleviation of this issue by immobilization adds extra costs associated with production time and additional materials. Whole-cell biocatalysis can also require complex downstream processing and is generally hampered by low substrate concentrations. To realize the full potential of biocatalysis, a long-term approach might instead harness nature’s tandem biocatalytic approach to the construction of complex secondary metabolites for the production of synthetic molecules.213 Whilst product concentration is generally lower than that of a chemical process, this is offset by the ability to generate molecules of high complexity in a single step and to eliminate costly isolation steps. Fermentation scientists have been harnessing natural, highly selective biosynthetic pathways to produce complex pharmaceutical intermediates from cheap raw materials for decades. Some of the most important pharmaceutical core molecules, such as penicillins and cephalosporins, are economically produced in this way. The wide differences between biosynthetic and chemical approaches to a target API can be gleaned by comparison of the alternative routes that have been reported for the synthesis of orlistat (()-tetrahydrolipstatin), a potent gastrointestinal lipase inhibitor used in the treatment of obesity (Figure 1.14). Orlistat can be prepared by hydrogenation of the highly lipophilic secondary metabolite lipstatin. Lipstatin itself is produced by fermentation (or, more correctly, a tandem biotransformation) from linoleic acid via a key enzyme-catalysed Claisen condensation using Streptomyces toxytricini under aerobic conditions.214 In contrast, the chemical approach to orlistat, based on the classical resolution or asymmetric synthesis of a highly functionalized six-membered ring lactone,215,216 is considered to be one of the most complex in the pharmaceutical industry, requiring four isolation steps and a number of protection/deprotections.217 However, molecules currently produced by fermentation are usually natural products, whereas most current drug candidates are synthetic. If lipstatin was not known to be a NHCHO

NHCHO O

O O

O

O

O O

O

H23C11 Lipstatin

Figure 1.14

Orlistat

Structures of lipstatin and orlistat

1.8 Concluding Remarks

69

natural product, would a biosynthetic approach have been developed? Most likely, biocatalytic approaches would be limited to the insertion of individual transformations into the current chemical route. In fact, lipase resolution of the six-membered lactone intermediate produced from the chemical approach to orlistat has been reported.218 Metabolic engineering of unnatural biosynthetic pathways, by the insertion of non-native genes into a host organism, offers great hope in this respect but is currently still in its infancy.214 The production of thymidine represents the first example of its successful implementation in pharmaceutical production (Scheme 1.23). Thymidine, although a synthetic molecule, bears considerable resemblance to other natural products, whereas many drug candidates have no counterpart in nature and will likely require transformations for which there is no biocatalytic precedent. To build an entirely artificial biosynthetic pathway using genetically modified organisms would require a monumental screening effort, given that the vast majority of enzymes involved in biosynthetic pathways have not yet been characterized and their specificities remain unevaluated. Furthermore, should it be necessary to insert a chemocatalytic step into the middle of a biosynthesis, transport across cell membranes would also require consideration. An alternative approach might instead be to express the required enzymes together in a genetically modified microorganism and use partially purified isolates, perhaps in tandem with chemocatalysts. The one-pot synthesis of corrin, a biosynthetic intermediate of vitamin B12, with 20 % unoptimized yield by 12 isolated enzymes demonstrates that complex tandem processes are feasible using isolated enzymes (Scheme 1.61),219 and the numerous chemoenzymatic processes available in the literature (some of which appear later in the book) demonstrate that chemocatalysts can be efficiently inserted into biocatalytic processes. HO2C

CO2H CO2H

HO2C

O H2N

N

12 Enzymes CO2H

H HO2C

N

HO2C

N H Vitamin B12

N

CO2H Corrin

Scheme 1.61 Tandem biocatalytic synthesis of corrin

1.8

Concluding Remarks

Biocatalysis has enjoyed widespread application in the preparation of chiral building blocks but has generally been employed on a limited basis for the production of more complex, late-stage pharmaceutical intermediates. Owing to pressure on the industry to develop more efficient and greener processes, along with rapid advances in the field of biocatalysis, this is beginning to change.220

70


The recent commercialization of more diverse ranges of enzymes, combined with a plethora of successful applications originating from both academia and the fine chemical industry, is placing biocatalysis in the mainstream as an addition to the chemist’s toolbox rather than an exotic curiosity. It is likely that, as the field matures, a greater diversity of non-natural molecules of greater complexity will become accessible through the tandem use of biocatalysts and genetically modified microorganisms. Together with advances in chemocatalysis, this will significantly impact on pharmaceutical production by improving efficiency and reducing waste.

Acknowledgements I would like to thank John Gray and Shiping Xie for their help in proof reading this chapter.

References 1. PricewaterhouseCoopers, Pharma 2020: The vision, 2007. http://www.pwc.com/Extweb/onlineforms.nsf/docid/DA9F87D21EE8BCAA852575A5005B3484?opendocument&doc¼vision. 2. McAndrews, P., Lilly sings a new tune: chorus unit brings high efficiency note to early R&D. The Pink Sheet, 2007, 69, 26. 3. Farina, V., Reeves, J.T., Senanayake, C.H. and Song, J.J., Asymmetric synthesis of active pharmaceutical ingredients. Chem. Rev., 2006, 106, 2734–2793. 4. Agrawal, Y.K., Bhatt, H.G., Raval, H.G., Oza, P.M. and Gogoi, P.J., Chirality – a new era of therapeutics. Mini-Rev Med. Chem., 2007, 7, 451–460. 5. KPMG, Pressure points: risk management in the pharmaceuticals industry, 2006. 6. US Food and Drug Administration, Pharmaceutical current good manufacturing practices (cGMPs) for the 21st century – a risk-based approach: final report, 2004. 7. McKenzie, P., Kiang, S., Tom, J., Rubin, A.E. and Futran, M., Can pharmaceutical process development become high tech? Am. Inst. Chem. Eng., 2006, 52, 3990–3994. 8. US Food and Drug Administration, Guidance for industry PAT – a framework for innovative pharmaceutical development, manufacturing, and quality assurance, 2004. 9. Rubin, A.E., Tummala, S., Both, D.A., Wang, C. and Delaney, E.J., Emerging technologies supporting chemical process R&D and their increasing impact on productivity in the pharmaceutical industry. Chem. Rev., 2006, 106, 2794–2810. 10. Straathof, A.J.J., Panke, S. and Schmid, A., The production of fine chemicals by biotransformations. Curr. Opin. Biotechnol., 2002, 13, 548–556. 11. Liese, A., Seelbach K. and Wandrey C. (eds), Industrial Biotransformations. Wiley–VCH, 2006. 12. Pollard, D.J. and Woodley, J.M., Biocatalysis for pharmaceutical intermediates: the future is now. Trends Biotechnol., 2007, 25, 66–73. 13. Panke, S., Held, M. and Wubbolts, M., Trends and innovations in industrial biocatalysis for the production of fine chemicals. Curr. Opin. Biotechnol., 2004, 15, 272–279. 14. Carey, J.S., Laffan, D., Thomson, C. and Williams, M.T., Analysis of the reactions used for the preparation of drug candidate molecules. Org. Biomol. Chem., 2006, 4, 2337–2347. 15. Breuer, M., Ditrich, K., Habicher, T., Hauer, B., Keßeler,M., Stu¨rmer,R. and Zelinski, T., Industrial methods for the production of optically active intermediates. Angew. Chem. Int. Ed., 2004, 43, 788 – 824. 16. Constable, D.J.C., Dunn, P.J., Hayler, J.D., Humphrey, G.R., Leazer Jr, J.L., Linderman, R.J., Lorenz, K., Manley, J., Pearlman, B.A., Wells, A., Zaks, A. and Zhang, T.Y., Key green chemistry research areas – a perspective from pharmaceutical manufacturers. Green Chem., 2007, 9, 411–420.

References

71

17. Schoemaker, H.E., Mink, D., Wubbolt, M.G., Dispelling the myths – biocatalysis in industrial synthesis. Science, 2003, 299, 1694–1697. 18. Vink, M.K.S. and Rozzell, J.D., Expanded opportunities for biocatalysis. PharmaChem, May 2006, 17–18. 19. (a) Lefevre, F., Jarrin, C., Ginolhac, A., Auriol, D. and Nalin, R., Environmental metagenomics: an innovative resource for industrial biocatalysis. Biocatal. Biotrans., 2007, 25, 242–250. (b) Ferrer, M., Martı`nez-Abarca,F. and Golyshin, P.N., Mining genomes and ‘metagenomes’ for novel catalysts. Curr. Opin. Biotechnol., 2005, 16, 588–593. (c) Cowan, D.A., Arslanoglu, A., Burton, S.G., Baker, G.C., Cameron, R.A., Smith, J.J. and Meyer, Q., Metagenomics, gene discovery and the ideal biocatalyst. Biochem. Soc. Trans., 2004, 32, 298–302. 20. For some recent reviews on enzyme modification see: (a) Reetz, M.T., Evolution in the test-tube as a means to creat selective biocatalysts. Chimia, 2007, 61, 100–103; (b) Sylvestre, J., Chautard, H., Cedrone, F. and Delcourt, M., Directed evolution of biocatalysts. Org. Proc. Res. Dev., 2006, 10, 562–571; (c) Hibbert, E.G. and Dalby, P.A., Directed evolution strategies for improved enzymatic performance. Microb. Cell Fact., 2004, 4, 29; (d) Otten, L.G. and Quax, W.J., Directed evolution: selecting today’s biocatalysts. Biomol. Eng., 2005, 22, 1–9. 21. For some recent reviews on HTS, see: (a) Aharoni, A., Griffiths, A.D. and Tawfik, D.S., Highthroughput screens and selections of enzyme-encoding genes. Curr. Opin. Chem. Biol., 2005, 9, 210–216; (b) Reymond, J. and Babiak, P., Screening systems. Adv. Biochem. Eng Biotechnol., 2007, 105, 31–58. 22. Fox, R.J., Davis, S.C., Mundorff, E.C., Newman, L.M., Gavrilovic, V., Ma, S.K., Chung, L.M., Ching, C., Tam, S., Muley, S., Grate, J., Gruber, J., Whitman, J.C., Sheldon, R.A. and Huisman, G.W., Improving catalytic function by ProSAR-driven enzyme evolution. Nat. Biotechnol., 2007, 25, 338–344. 23. Hilterhaus, L. and Liese, A., Building blocks. Adv. Biochem. Eng. Biotechnol., 2007, 105, 133–173. 24. Grossman, S.J., Overview: drug metabolism in the modern pharmaceutical industry. In Drug Metabolism in Drug Design and Development, Zhang, D., Zhu, M. and Humphreys, W.G. (eds). J. Wiley & Sons, Inc., 2008, pp. 3–13. 25. Testa, B. and Kra¨mer, S. D., The biochemistry of drug metabolism – an introduction part 1. Principles and overview. Chem. Biodiv., 2006, 3, 1053–1101 and references cited therein. 26. Azerad, R., Microbial models for drug metabolism. Adv. Biochem. Eng. Biotechnol., 1999, 63, 169–218. 27. Smith, R.V. and Rosazza, J.P., Microbial models of mammalian metabolism. J. Pharm. Sci., 1975, 64, 1737–1759. 28. Alexandre, V., Ladril, S., Maurs, M. and Azerad, R., Microbial models of animal drug metabolism Part 5. Microbial preparation of human hydroxylated metabolites of irbesartan. J. Mol. Catal. B: Enzymatic, 2004, 29, 173–179. 29. For a recent review on microbial preparation of metabolites, see Venisetty, R.K. and Ciddi, V., Application of microbial biotransformation for the new drug discovery using natural drugs as substrates. Curr. Pharm. Biotechnol., 2003, 4, 153–167. 30. Testa, B. and Kra¨mer, S.D., The biochemistry of drug metabolism – an introduction part 2. Redox reactions and their enzymes. Chem. Biodiv., 2007, 4, 257–404. 31. Smith, D.A., Ackland, M.J. and Jones, B.C., Properties of cytochrome P450 isoenzymes and their substrates. Part 2: properties of cytochrome P450 substrates. Drug Discov. Today, 1997, 2, 479–486 and references cited therein. 32. (a) Hanlon, S.P., Friedburg, T., Wolf, C.R., Ghisalba, O. and Kittelmann, M., Recombinant yeast and bacteria that express human P450s: bioreactors for drug discovery, development, and biotechnology. In Modern Biooxidation. Enzymes, Reactions and Applications. Schmid, R.D. and Urlacher, V.B. (eds). Wiley–VCH: Weinheim, 2007, pp. 233–252; (b) Crespi, C.L. and Miller, V.P., The use of heterologously expressed drug metabolizing enzymes – state of the art and prospects for the future. Pharmacol. Therap., 1999, 84, 121–131. 33. Schwaneberg, U., Otey, C., Cirino, P.C., Farinos, E. and Arnold, F.H., Cost-effective whole-cell assay for laboratory evolution of hydroxylases in Escherichia coli. J. Biomol. Screen., 2001, 6, 111–117.

72


34. Ghisalba, O. and Kittelmann, M., Preparation of drug metabolites using fungal and bacterial strains. In Modern Biooxidation, Schmid, R.D. and Urlacher, V. (eds). Wiley–VCH: Weinheim, 2007, pp. 211–232. 35. This volume, Section 12.3, Kittelmann et al. 36. Wong, J.W., Watson Jr, H.A., Bouressa, J.F., Burns, M.P., Cawley, J.J., Doro, A.E., Guzek, D.B., Hintz, M.A., McCormick, E.L., Scully, D.A., Siderewicz, J.M., Taylor, W.J., Truesdell, S.J. and Wax, R.G. Biocatalytic oxidation of 2-methylquinoxaline to 2-quinoxalinecarboxylic acid. Org. Proc. Res. Dev., 2002, 6, 477–481. 37. Stachulski, A.V. and Jenkins, G.N., The synthesis of O-glucuronides. Nat. Prod. Rep., 1998, 15, 173–186. 38. Chiu, S.H. and Huskey, S.W., Species differences in N-glucuronidation. Drug Metab. Dispos., 1998, 26, 838–847. 39. Kittelmann, M., Rheinegger, U., Espigat, A., Oberer, L., Aichholz, R., Francotte, E. and Ghisalba, O., Preparative enzymatic synthesis of the acylglucuronide of mycophenolic acid. Adv. Synth. Catal., 2003, 345, 825–829. 40. Baratto, L., Candido, A., Marzorati, M., Sagui, F., Riva, S. and Danieli, B., Laccase-mediated oxidation of natural glycosides. J. Mol. Catal. B: Enzymatic, 2006, 39, 3–8. 41. (a) Fabbrini, M., Galli, C., Gentili, P. and Macchitella, D., An oxidation of alcohols by oxygen with the enzyme laccase and mediation by TEMPO. Tetrahedron Lett., 2001, 42, 7551–7553; (b) NikuPaavola, M.-L. and Viikari, L., Enzymatic oxidation of alkenes. J. Mol. Catal. B: Enzymatic, 2000, 10, 435–444; (c) Potthast, A., Rosenau, T., Chen,C.-L. and Gratzl, J.S., Selective enzymatic oxidation of aromatic methyl groups to aldehydes. J. Org. Chem. 1995, 60, 4320–4321. 42. Wong, C.-H. and Whitesides, G.M., Enzymes in synthetic organic chemistry. In Tetrahedron Organic Chemistry Series, Baldwin, J.E. and Magnus, P.D. (eds), Pergamon, 1994. 43. Newman, D.J., Cragg, G.M. and O’Keefe, B.R., Biopharmaceutical drugs from natural sources. In Modern Biopharmaceuticals, Kna¨blein, J. (ed). Wiley–VCH: Weinheim, 2005, pp. 451–496. 44. Walsh, G. and Jefferis, R., Post-translational modifications in the context of therapeutic proteins. Nat. Biotechnol., 2006, 24, 1241–1252. 45. Meyer, A., Pellaux, R. and Panke, S. Bioengineering novel in vitro metabolic pathways using synthetic biology. Curr. Opin. Microbiol., 2007, 10, 246–253. 46. (a) Koeller, K.M. and Wong, C.-H., Synthesis of complex carbohydrates and glycoconjugates: enzyme-based and programmable one-pot strategies. Chem. Rev. 2000, 100, 4465–4493; (b) Khmelnitsky, Y.L., Current strategies for in vitro protein glycosylation. J. Mol. Catal. B, 2004, 31, 73–81. ¨ hrlein, R., Glycosyltransferase-catalysed synthesis of non-natural oligosaccharides. Topics 47. O Curr. Chem., 1999, 200, 227–254. 48. Rowan, A.S. and Hamilton, C.J., Recent developments in preparative enzymatic syntheses of carbohydrates. Nat. Prod. Rep., 2006, 23, 412–443 and references cited therein. 49. Ullman, C.G. and Perkins, S.J., A classification of nucleotide diphospho-sugar glycosyltransferases based on amino acid sequence similarities. Biochem. J., 1997, 326, 929–942. 50. Shao, H., He, X., Achnine, L., Blount, J.W., Dixon, R.A. and Wang, X., Crystal structures of a multifunctional triterpene/flavonoid glycosyltransferase from Medicago truncatula. The Plant Cell, 2005, 17, 3141–3154. 51. Lim, E.-K., Ashford, D.A., Hou, B., Jackson, R.G. and Bowles, D.J., Biotechnol. Bioeng., 2004, 87, 623–631. 52. Perusino, G., Cobucci-Ponzano, B., Rossi, M. and Moracci, M., Recent advances in the oligosaccharide synthesis promoted by catalytically engineered glycosidases. Adv. Synth. Catal., 2005, 347, 941–950. 53. Mackenzie, L.F., Wang, Q., Warren, R.A.J. and Withers, S.G., Glycosynthases: mutant glycosidases for oligosaccharide synthesis. J. Am. Chem. Soc., 1998, 120, 5583–5584. 54. Spence, D.W. and Ramsden, M., Penicillin acylases. In Industrial Enzymes, Polaina, J. and MacCabe, A.P. (eds). Springer: Dordrecht, 2007, pp. 583–597. 55. Henderson R.K., Jimeńez-Gonza´lez C., Preston C., Constable D.J.C. and Woodley, J.M., EHS & LCA assessment for 7-ACA synthesis. A case study for comparing biocatalytic & chemical synthesis. Ind. Biotechnol., 2008, 4, 180–192.

References

73

56. Fildes, R.A., Potts, J.R. and Farthing, J.E., Process for preparing cephalosporin derivatives. PCT Appl., 1974, US 3,801,458. 57. Matsuda, A., Matsuyama, K., Yamamoto, K., Ichikawa, S. and Komatsu, K.-I., Cloning and characterization of the genes for two distinct cephalosporin acylases from a Pseudomonas strain. J. Bacteriol., 1987, 169, 5815–5820. 58. Matsumoto, K., Production of 6-APA, 7-ACA and 7-ADCA by immobilised penicillin and cephalosporin amidases. Bioproc. Technol. 1993, 16, 67–88. 59. Brakhage, A.A., Molecular regulation of b-lactam biosynthesis in filamentous fungi. Microbiol. Mol. Biol. Rev., 1998, 62, 547–585. 60. Conder, M.J., Crawford, L., McAda, P.C. and Rambosek, J.A., Novel bioprocess for preparing 7-ADCA. PCT Appl., 1992, EP 0532341 A1. 61. Sheldon, R.A. and van Rantwijk, F., Biocatalysis for sustainable organic synthesis. Aust. J. Chem., 2004, 57, 281–289. 62. Fernańdez-Lafuente,R., Rosell, C.M., Piatkowska, B. and Guisań, J.M., Synthesis of antibiotics (cephaloglycin) catalyzed by penicillin G acylase: evaluation and optimization of different synthetic approaches. Enzyme Microb. Technol., 1996, 19, 9–14. 63. Kallenberg, A.I., van Rantwijk, F. and Sheldon, R.A., Immobilization of penicillin G acylase: the key to optimum performance. Adv. Synth. Catal., 2005, 347, 905–926. 64. Schroen, C.G.P.H., Nierstrasz, V.A., Bosma, R., Kroon, P.J., Tjeerdsma, P.S., DeVroom, E., VanderLaan, J.M., Moody, H.M., Beeftink, H.H., Janssen, A.E.M. and Tramper, J., Integrated reactor concepts for the enzymatic kinetic synthesis of cephalexin. Biotech. Bioeng., 2002, 80, 144–155. 65. Kim, I., Song, X., Vig, B.S., Mittal, S., Shin, H.-C., Lorenzi, P.J. and Amidon, G.L., A novel nucleoside prodrug-activating enzyme: substrate specificity of biphenyl hydrolase-like protein. Mol. Pharm., 2004, 1, 117–127. 66. Tamarez, M., Morgan, B., Wong, G.S.K., Tong, W., Bennett, F., Lovey, R., McCormick, J.L. and Zaks, A., Pilot-scale lipase-catalyzed regioselective acylation of ribavirin in anhydrous media in the synthesis of a novel prodrug intermediate. Org. Proc. Res. Dev., 2003, 7, 951–953. 67. (a) Zaks, A. and Klibanov, A.M., Enzymatic catalysis in nonaqueous solvent. J. Biol. Chem., 1988, 263, 3194–3201; (b) Klibanov, A.M., Asymmetric enzymatic oxidoreductions in organic solvents. Curr. Opin. Biotechnol., 2003, 14, 427–431. 68. Ferrero, M. and Gotor, V., Biocatalytic selective modifications of conventional nucleosides, carbocyclic nucleosides, and C-nucleosides. Chem. Rev., 2000, 100, 4319–4347 and references cited therein. 69. Tamarez, M., Morgan, B., Wong, G.S.K., Tong, W., Bennett, F., Lovey, R., McCormick, J.L. and Zaks, A., Pilot-scale lipase-catalyzed regioselective acylation of ribavirin in anhydrous media in the synthesis of a novel prodrug intermediate. Org. Proc. Res. Dev., 2003, 7, 951–953. 70. Mahmoudian, M., Eaddy, J. and Dawson, M., Enzymic acylation of 506U78 (2-amino-9--Darabinofuranosyl-6-methoxy-9H-purine), a powerful new anti-leukaemic agent. Biotechnol. Appl. Biochem., 1999, 29, 229–233. 71. Hanson, R.L., Shi, Z., Brzozowski, D.B., Banerjee, A., Kissick, T.P., Singh, J., Pullockaran, A.J., North, J.T., Fan, J., Howell, J., Durand, S.C., Montana, M.A., Kronenthal, D.R., Mueller, R.H. and Patel, R.N., Regioselective enzymatic aminoacylation of lobucavir to give an intermediate for lobucavir prodrug. Bioorg. Med. Chem., 2000, 8, 2681–2687. 72. Manzoni, M. and Rollini, M., Biosynthesis and biotechnological production of statins by filamentous fungi and application of these cholesterol-lowering drugs. Appl. Microbiol. Biotechnol., 2002, 58, 555–564. 73. Sleteinger, M., Verhoeven, T.R. and Volante, R.P., Process for C-methylation of 2-methylbutyrates. US PCT Appl., 1986, US 4,582,915. 74. Askin, D., Verhoeven, T.R., Liu, T.M.-H. and Shinkai, I., Synthesis of synvinolin: extremely high conversion alkylation of an ester enolate. J. Org. Chem., 1991, 56, 4929–4932. 75. Schimmel, T.G., Borneman,W.S. and Conder, M.J., Purification and characterization of a lovastatin esterase from Clonostachys compactiuscula. Appl. Environ. Microbiol., 1997, 63, 1307–1311.

74


76. Xie, X. and Tang, Y., Efficient synthesis of simvastatin by use of whole-cell biocatalysis. Appl. Environ. Microbiol., 2007, 73, 2054–2060. 77. Xie, X., Wong, W.W. and Tang, Y., Improving simvastatin bioconversion in Escherichia coli by deletion of bioH. Metabol. Eng., 2007, 9, 379–386. 78. Kern, A., Tilley, E., Hunter, I.S., Legıˇsa, M. and Glieder, A., Engineering primary metabolic pathways of industrial micro-organisms. J. Biotechnol., 2007, 129, 6–29. 79. Sørensen, H.P. and Mortensen, K.K. Advanced genetic strategies for recombinant protein expression in Escherichia coli. J. Biotechnol., 2005, 115, 113–128. 80. McCandliss, R.J. and Anderson, D.M., Fermentation process for the production of pyrimidine deoxyribonucleosides. PCT Int. Appl., 1991, WO91/09130. 81. Anderson, D. M., Liu, L., Podkovyrov, S. and Wang, B., PCT Int. Appl., 2001, WO2001002580 A1. 82. Anderson, D.M., Collis, A.J., Liu, L., Podkovyrov, S. and Preston, C., Thymidine production in various strains/constructs. PCT Int. Appl., 2007, WO2007/090810 A1. 83. Roth, B.D., The discovery and development of atorvastatin, a potent novel hypolipidemic agent. Prog. Med. Chem., 2002, 40, 1–22 and references cited therein. 84. Mu¨ller, M., Chemoenzymatic synthesis of building blocks for statin side chains. Angew. Chem. Int. Ed., 2005, 44, 362–365. 85. Fox, R.J., Davis, S.C., Mundorff, E.C., Newman, L.M., Gavrilovic, V., Ma, S.K., Chung, L.M., Ching, C., Tam, S., Muley, S., Grate, J., Gruber, J., Whitman, J.C., Sheldon, R.A. and Huisman, G.W., Improving catalytic function by ProSAR-driven enzyme evolution. Nat. Biotechnol., 2007, 25, 338–344. 86. Lewkowicz, E.S. and Iribarren, A.M., Nucleoside phosphorylases. Curr. Org. Chem., 2006, 10, 1197–1215; Utagawa, T., Enzymatic preparation of nucleoside antibiotics. J. Mol. Catal. B, 1999, 6, 215–222. 87. Shirae, H., Yokozeki, K., Uchiyama, M. and Kubota, K., Enzymatic production of ribavirin from purine nucleosides by Brevibacterium acetylicum. Agric. Biol. Chem., 1988, 52, 1777– 1783. 88. Krenitsky, T.A., Koszalka, G.W. and Tuttle, J.V., Purine nucleoside synthesis, an efficient method employing nucleoside phosphorylases. Biochemistry, 1981, 20, 3615–3621. 89. Krenitsky, T.A., Elion, G.B. and Rideout, J.L., Enzymatic synthesis of arabinonucleosides, Eur. PCT Appl., 1979, EP 0002192; Krenitsky, T.A., Koszalka, G.W., Tuttle, J.V., Rideout, J.L. and Elion, G.B., An enzymatic synthesis of purine D-arabinonucleosides. Carbohydrate Res., 1981, 97, 139–146. 90. Komura, H., Yoshino, T. and Ishido, Y., Synthetic studies by the use of carbonates. II. Easy method of preparing cyclic carbonates of polyhydroxy compounds by transesterification with ethylene carbonate. Bull. Chem. Soc., 1973, 46, 550–553. 91. Averett, D.R., Koszalka, G.W., Fyfe, J.A., Roberts, G.B., Purifoy, D.J.M. and Krenitsky, T.A., 6-Methoxypurine arabinoside as a selective and potent inhibitor of varicella-zoster virus. Antimicrob. Agents Chemother. 1991, 35, 851–857. 92. Mahmoudian, M., Development of bioprocesses for the generation of anti-inflammatory, antiviral and anti-leukaemic agents. Focus Biotechnol. 2001, 1, 249–265. 93. Utagawa, T., Morisawa, H., Yoshinaga, F., Yamazaki, A., Misugi, K. and Hirose, Y., Microbiological synthesis of adenine arabinoside. Agric. Biol. Chem., 1985, 49, 1053–1058. 94. Herbal, K., Kitteringham, J., Voyle, M. and Whitehead, A.J., Synthesis of the enantiomer of nelarabine. Tetrahedron Lett., 2005, 46, 2961–2964. 95. Warren, L. and Felsenfeld, H., Biosynthesis of N-acetylneuraminic acid. Biochem. Biophys. Res. Commun., 1961, 4, 232–235. 96. Tabata, K., Koizumi, S., Endo, T. and Ozaki, A., Production of N-acetyl-D-neuraminic acid by coupling bacteria expressing N-acetyl-D-glucosamine 2-epimerase and N-acetyl-D-neuraminic acid synthetase. Enzyme Microb. Technol. 2002, 30, 327–333. 97. Samain, E., High yield production of sialic acid (Neu5Ac) by fermentation. PCT Int. Appl., 2008, WO 2008040717 A2; Lundgren, B.R. and Boddy, C.N., Sialic acid and N-acyl sialic acid analog production by fermentation of metabolically and genetically engineered Escherichia coli. Org. Biomol. Chem., 2007, 5, 1903–1909.

References

75

98. Comb, D.G. and Roseman, S., Composition and enzymic synthesis of N-acetylneuraminic acid (sialic acid). J. Am. Chem. Soc., 1958, 80, 497–499. 99. Cipolletti, G., Tamerlani, G., Lombardi, I. and Bartalucci, D., Process for the preparation of N-acetyl-D-mannosamine monohydrate. PCT Int. Appl., 2007, WO 2007135086 A1. 100. Mahmoudian, M., Noble, D., Drake, C.S., Middleton, R.F., Montgomery, D.S., Piercey, J.E., Ramlakhan, D., Todd, M. and Dawson, M.J., An efficient process for production of N-acetylneuraminic acid using N-acetylneuraminic acid aldolase. Enzyme Microb. Technol. 1997, 20, 393–400. 101. Kragl, U., Wandrey, C., Ghisalba, O. and Gygax, D., Enzymic preparation of N-acetyl neuraminic acid. Ger. Offen. 1991, DE 3937891 A1; Maru, I., Ohnishi, J., Ohta, Y. and Tsukada, Y., Why is sialic acid attracting interest now? Complete enzymatic synthesis of sialic acid with N-acylglucosamine 2-epimerase. J. Biosci. Bioeng., 2002, 93, 258–265; Maru, I., Ohnishi, J., Ohta, Y. and Tsukada, Y., Simple and large-scale production of N-acetylneuraminic acid from N-acetyl-D-glucosamine and pyruvate using N-acetyl-D-glucosamine 2-epimerase and N-acetylneuraminate lyase. Carbohydrate Res., 1998, 306, 575–578. 102. Kragl, U., Kittelmann, M., Ghisalba, O. and Wandrey, C., N-Acetylneuramic acid: from a rare chemical from natural sources to a multikilogram enzymatic synthesis for industrial application. Ann. N. Y. Acad. Sci., 1995, 750, 300–305. 103. Garcıà-Urdiales, E., Alfonso, I. and Gotor, V., Enantioselective enzymatic desymmetrizations in organic synthesis. Chem. Rev., 2005, 105, 313–354. 104. (a) Hasan, F., Shah, A.A. and Hameed, A. Industrial applications of microbial lipases. Enzyme Microb. Technol., 2006, 39, 235–251; (b) Maurer, K.-H., Detergent proteases. Curr. Opin. Biotechnol., 2004, 15, 330–334; (c) Gupta, R., Beg, Q.K. and Lorenz, P., Bacterial alkaline proteases: molecular approaches and industrial applications. Appl. Microbiol. Biotechnol., 2002, 59, 15–32. 105. Martinez, C.A., Hu, S., Dumond, Y., Tao, J., Kelleher, P. and Tully, L., Development of a chemoenzymatic manufacturing process for pregabalin. Org. Proc. Res. Dev., 2008, 12, 392–398. 106. Canoy, W.L., Cooley, B.E., Corona, J.A., Lovelace, T.C., Millar, A., Weber, A.M., Xie, S. and Zhang, Y., Efficient synthesis of (3R,3aS,6aR)-hexahydrofuro[2,3-b]furan-3-ol from glycolaldehyde. Org. Lett., 2008, 10, 1103–1106 and references cited therein. 107. Yu, R.H., Polniaszek, R.P., Becker, M.W., Cook, C.M. and Yu, L.H.L., Research and development of an efficient synthesis of hexahydrofuro[2,3-b]furan-3-ol moiety – a key component of the HIV protease inhibitor candidates. Org. Proc. Res. Dev., 2007, 11, 972–980. 108. Doan, B.D., Davis, R.D. and Lovelace, T.C., Process for preparing intermediates for HIV aspartyl protease inhibitors, particularly (3R,3aS,6aR)-hexahydrofuro[2,3-b]furan-3-ol and its (3R,3aS,6aR)-enantiomer. PCT Int. Appl., 2003, WO 2003024974 A2. 109. Roberts, J., A commercially viable route and process to HIV PI GW640385X. Abstracts of Papers, 226th ACS National Meeting, New York, NY, United States, 2003, September 7–11. 110. Vaidyanathan, R., Hesmondhalgh, L. and Hu, S., A chemoenzymatic synthesis of an androgen receptor antagonist. Org. Proc. Res. Dev., 2007, 11, 903–906. 111. Terao, Y., Tsuji, K., Murata, M., Achiwa, K., Nishio, T., Watanabe, N. and Seto, K., Facile process for enzymatic resolution of racemic alcohols. Chem. Pharm. Bull., 1989, 37, 1653–1655. 112. Andrews, I.P., Atkins, R.J., Breen, G.F., Carey, J.S., Forth, M.A., Morgan, D.O., Shamji, A., Share, A.C., Smith, S.A.C., Walsgrove, T.C. and Wells, A.S., The development of a manufacturing route for the GPIIb/IIIa receptor antagonist SB-214857-A. Part 1: synthesis of the key intermediate 2,3,4,5-tetrahydro-4-methyl-3-oxo-1H-1,4-benzodiazepine-2-acetic acid methyl ester, SB-235349. Org. Proc. Res. Dev., 2003, 7, 655–662; Atkins, R.J., Banks, A., Bellingham, R.K., Breen, G.F., Carey, J.S., Etridge, S.K., Hayes, J.F., Hussain, N., Morgan, D.O., Oxley, P., Passey, S.C., Walsgrove, T.C. and Wells, A.S., The development of a manufacturing route for the GPIIb/IIIa receptor antagonist SB-214857-A. Part 2: conversion of the key intermediate SB-235349 to SB-214857-A. Org. Proc. Res. Dev., 2003, 7, 663–675. 113. Roberts, N.J., Seago, A., Carey, J.S., Freer, R., Preston, C. and Lye, G.J., Lipase catalysed resolution of the lotrafiban intermediate 2,3,4,5-tetrahydro-4-methyl-3-oxo-1H-1,4-benzodiazepine-2-acetic acid methyl ester in ionic liquids: comparison to the industrial t-butanol process. Green Chem., 2004, 6, 475–482.

76


114. Butters, M., Catterick, D., Craig, A., Curzons, A., Dale, D., Gillmore, A., Green, S.P., Marziano, I., Sherlock, J.-P. and White, W., Critical assessment of pharmaceutical processess – a rationale for changing the synthetic route. Chem. Rev., 2006, 106, 3002–3027. 115. Casu, F., Chiacchio, M.A., Romeo, R. and Gumina, G., Chiral synthesis of heterosubstituted nucleoside analogs from non-carbohydrate precursors. Curr. Org. Chem., 2007, 11, 1017– 1032. 116. Mahmoudian, M., Baines, B.S., Drake, C.S., Hale, R.S., Jones, P., Piercey, J.E., Montgomery, D.S., Purvis, I.J., Storer, R., Dawson, M.J. and Lawrence, G.C., Enzymatic production of optically pure (20 R-cis)-20 -deoxy-30 -thiacytidine (3TC, lamivudine): a potent anti-HIV agent. Enzyme Microb. Technol., 1993, 15, 749–755. 117. Munch-Petersen, A., Nygaard, P., Hammer-Jespersen, K. and Fiil, N., Mutants constitutive for nucleoside-catabolizing enzymes in Escherichia coli K 12. Eur. J. Biochem., 1972, 27, 208–215. 118. Mahmoudian, M. and Dawson, M.J., Chemoenzymic production of the antiviral agent EpivirTM. In Biotechnology of Antibiotics, Strohl, W.R. (ed.). Dekker: New York, 1997, pp. 753–777. 119. Solares, L.F., Brieva, R., Quiro´s, M., Llorente, I., Bayod, M. and Gotor, V., Enzymatic resolution of a quaternary stereogenic centre as the key step in the synthesis of (S)-(þ)citalopram. Tetrahedron Asymm., 2004, 15, 341–345. 120. Mateja Pogorevc, M. and Faber, K., Biocatalytic resolution of sterically hindered alcohols, carboxylic acids and esters containing fully substituted chiral centers by hydrolytic enzymes. J. Mol. Catal. B, 2000, 10, 357–376 and references cited therein. 121. Taylor, S.J.C., Sutherland, A.G., Lee, C., Wisdom, R., Thomas, S., Roberts, S.M. and Evans, C., Chemoenzymatic synthesis of ()-carbovir utilizing a whole cell catalysed resolution of 2-azabicyclo[2.2.1 ]hept-5-en-3-one. J. Chem. Soc. Chem. Commun., 1990, 1120–1121; Evans, C.T., Roberts, S.M., Shoberu, K.A. and Sutherland, A.G., Potential use of carbocyclic nucleosides for the treatment of AIDS: chemo-enzymatic syntheses of the enantiomers of carbovir. J. Chem. Soc. Perkin Trans. 1, 1992, 589–592. 122. Taylor, S.J.C, McCague, R., Wisdom, R., Lee, C., Dickson, K., Ruecroft, G., O’Brien, F., Littlechild, J., Bevan, J., Roberts, S.M. and Evans, C.T., Development of the biocatalytic resolution of 2-azabicyclo [2,2,1] hept-5-en-3-one as an entry to single-enantiomer carbocyclic nucleosides. Tetrahedron Asymm., 1993, 4, 1117–1128. 123. Mahmoudian, M., Lowdon, A., Jones, M., Dawson, M. and Wallis, C., A practical enzymatic procedure for the resolution of N-substituted 2-azabicyclo[2.2.1]hept-5-en-3-one. Tetrahedron Asymm. 1999, 10, 1201–1206. 124. Carnell, A.J., Desymmetrisation of prochiral ketones using lipases. J. Mol. Catal. B, 2002, 19– 20, 83–92. 125. Allan, G., Carnell, A.J., Escudero Hernandez, M.L. and Pettman, A., Chemoenzymatic synthesis of a tachykinin NK-2 antagonist. Tetrahedron, 2001, 57, 8193–8202. 126. Carnell, A.J., Barkely, J. and Singh, A., Desymmetrisation of prochiral ketones by catalytic enantioselective hydrolysis of their enol esters using enzymes. Tetrahedron Lett., 1997, 38, 7781–7784; Allan, G.R., Carnell, A.J. and Kroutil, W., One-pot deracemisation of an enol acetate derived from a prochiral cyclohexanone. Tetrahedron Lett., 2001, 42, 5959–5962. 127. Pellissier, H., Recent developments in dynamic kinetic resolution. Tetrahedron, 2008, 64, 1563–1601; Turner, N.J., Enzyme catalysed deracemisation and dynamic kinetic resolution reactions. Curr. Opin. Chem. Biol., 2004, 8, 114–119; Gruber, C.C., Lavandera, I., Faber, K. and Kroutil, W., From a racemate to a single enantiomer: deracemisation by stereoinversion. Adv. Synth. Catal., 2006, 348, 1789–1805; Pellissier, H., Dynamic kinetic resolution. Tetrahedron, 2003, 59, 8291–8327; Pa`mies, O. and Ba¨ckvall, J.-E., Combination of enzymes and metal catalysts. A powerful approach in asymmetric catalysis. Chem. Rev., 2003, 103, 3247–3261. 128. Hughes, G., O’Shea, P.D., Devine, P.N., Foster, B., Gauthier, D., Limanto, J., Truppo, M., Pollard, D., Naber, J., McKay, D.J. and Volante, R.P., The discovery and development of odanacatib: a selective inhibitor of cathepsin K for the treatment of osteoporosis. In 25th SCI Process Development Symposium, 2007.

References

77

129. Gu, R.-L., Lee, I.S. and Sih, C.J., Chemo-enzymatic asymmetric synthesis of amino acids. Enantioselective hydrolyses of 2-phenyl-oxazolin-5-ones. Tetrahedron Lett., 1992, 33, 1953– 1956; Crich, J., Brieva, R., Marquart, P., Gu, R.-L., Flemming, S. and Sih, C.J., Enzymic asymmetric synthesis of -amino acids. Enantioselective cleavage of 4-substituted oxazolin-5ones and thiazolin-5-ones. J. Org. Chem., 1993, 58, 3252–3258. 130. Limanto, J., Shaifee, A., Devine, P.N., Upadhyay, V., Desmond, R.A., Foster, B.R., Gauthier Jr, D.R., Reamer, R.A. and Volante, R.P., An efficient chemoenzymatic approach to (S)-fluoroleucine ethyl ester. J. Org. Chem., 2005, 70, 2372–2375. 131. Madsen, R., Roberts, C. and Fraser-Reid, B., The pent-4-enoyl group: a novel amine-protecting group that is readily cleaved under mild conditions. J. Org. Chem., 1995, 60, 7920–7926. 132. Truppo, M.D. and Moore, J.C., Process for making fluoroleucine ethyl esters. US PCT Appl., 2007, US 2007/0059812 A1. 133. Pesti, J.A., Yin, J., Zhang, L.-H. and Anzalone, L., Reversible Michael reaction–enzymatic hydrolysis: a new variant of dynamic resolution. J. Am. Chem. Soc., 2001, 123, 11075–11076; Pesti, J.A., Yin, J., Zhang, L.-H., Anzalone, L., Waltermire, R.E., Ma, P., Gorko, E., Confalone, P.N., Fortunak, J., Silverman, C., Blackwell, J., Chung, J.C., Hrytsak, M.D., Cooke, M., Powell, L. and Ray, C., Efficient preparation of a key intermediate in the synthesis of roxifiban by enzymatic dynamic kinetic resolution on large scale. Org. Proc. Res. Dev., 2004, 8, 22–27. 134. Miyazawa, T., Kurita, S., Ueji, S., Yamada, T. and Shigeru, K., Resolution of mandelic acids by lipase-catalyzed transesterifications in organic media: inversion of enantioselectivity mediated by the acyl donor. J. Chem. Soc. Perkin Trans. 1, 1992, 18, 2253–2255. 135. Gro¨ger, H., Enzymatic routes to enantiomerically pure aromatic -hydroxy carboxylic acids: a further example for the diversity of biocatalysis. Adv. Synth. Catal., 2001, 343, 547–558. 136. Huang, H.-R., Xu, J.-H., Xu, Y., Pan, J. and Liu, X., Preparation of (S)-mandelic acids by enantioselective degradation of racemates with a new isolate Pseudomonas putida ECU1009. Tetrahedron Asymm., 2005, 16, 2113–2117. 137. DeSantis, G., Zhu, Z., Greenberg, W.A., Wong, K., Chaplin, J., Hanson, S.R., Farwell, B., Nicholson, L.W., Rand, C.L., Weiner, D.P., Robertson, D.E. and Burk, M.J., An enzyme library approach to biocatalysis: development of nitrilases for enantioselective production of carboxylic acid derivatives. J. Am. Chem. Soc., 2002, 124, 9024–9025. 138. Saksena, A.K., Girijavallabhan, V.M., Lovey, R.G., Pike, R.E., Desai, J.A., Ganguly, A.K., Hare, R.S., Loebenberg, D., Cacciapuoti, A. and Parmegiani, R.M., Enantioselective synthesis of the optical isomers of broad-spectrum orally active antifungal azoles, SCH 42538 and SCH 45012. Biorg. Med. Chem. Lett., 1994, 4, 2023–2028. 139. Morgan, B., Dodds, D.R., Zaks, A., Andrews, D.R. and Klesse, R., Enzymatic desymmetrisation of prochiral 2-substituted-1,3-propanediols: a practical chemoenzymatic synthesis of a key precursor of SCH51048, a broad-spectrum orally active antifungal agent. J. Org. Chem., 1997, 62, 7736–7743. 140. Schmid, R.D. and Verger, R., Lipases: interfacial enzymes with attractive applications. Angew. Chem. Int. Ed., 1998, 37, 1608–1633 and references cited therein. 141. Homann M.J., Suen W.-C., Zhang, N. and Zaks, A., Comparative analysis of chemical and biocatalytic syntheses of drug intermediates. In Biocatalysis in the Pharmaceutical and Biotechnology Industries, Patel, R.N. (ed.), CRC Press, 2007, pp. 645–659 and references cited therein. 142. Maddrell, S.J., Turner, N.J., Kerridge, A., Willetts, A.J. and Crosby, J., Nitrile hydratase enzymes in organic synthesis: enantioselective synthesis of the lactone moiety of the mevinic acids. Tetrahedron Lett., 1996, 37, 6001–6004. 143. Bergeron, S., Chaplin, D.A., Edwards, J.H., Ellis, B.S.W., Hill, C.L., Holt-Tiffin, K., Knight, J.R., Mahoney, T., Osborne, A.P. and Ruecroft, G., Nitrilase-catalysed desymmetrisation of 3hydroxyglutaronitrile: preparation of a statin side-chain intermediate. Org. Proc. Res. Dev., 2006, 10, 661–665. 144. DeSantis, G., Wong, K., Farwell, B., Chatman, K., Zhu, Z., Tomlinson, G., Huang, H., Tan, X., Bibbs, L., Chen, P., Kretz, K. and Burk, M.J., Creation of a productive, highly enantioselective nitrilase through gene site saturation mutagenesis (GSSM). J. Am. Chem. Soc., 2003, 125, 11476–11477.

78


145. Homann, M.J., Vail, R.B., Previte, E., Tamarez, M., Morgan, B., Dodds, D.R. and Zaks, A., Rapid identification of enantioselective ketone reductions using targeted microbial libraries. Tetrahedron, 2004, 60, 789–797. 146. Barbieri, C., Caruso, E., D’Arrigo, P., Fantoni, G.P. and Servi, S., Chemo-enzymatic synthesis of (R)- and (S)-3,4-dichlorophenylbutanolide intermediate in the synthesis of sertraline. Tetrahedron Asymm. 1999, 10, 3931–3937. 147. Leonida, M.D., Redox enzymes used in chiral syntheses coupled to coenzyme regeneration. Curr. Med. Chem., 2001, 8, 345–369. 148. Eckstein, M., Daubmann, T. and Kragl, U., Recent developments in NAD(P)H regeneration for enzymatic reductions in one- and two-phase systems. Biocatal. Biotrans., 2004, 22, 89–96. 149. Davis, S.C., Grate, J.H., Gray, D.R., Gruber, J.M., Huisman, G.W., Ma, S.K., Newman, L.M., Sheldon, R. and Wang, L.A., Enzymatic processes for the production of 4-substituted 3-hydroxybutyric acid derivatives. PCT Int. Appl., 2004, WO 2004015132 A2. 150. Ema, T., Okita, N., Ide, S. and Sakai, T., Highly enantioselective and efficient synthesis of methyl (R)-o-chloromandelate with recombinant E. coli: toward practical and green access to clopidogrel. Org. Biomol. Chem., 2007, 5, 1175–1176. 151. King, A.O., Corely, E.G., Anderson, R.K., Larsen, R.D., Verhoeven, T.R., Reider, P.J., Xiang, Y.B., Belley, M., Leblane, Y., Labelle, M., Prasit, P. and Zamboni, R.J., An efficient synthesis of LTD4 antagonist L-699,392. J. Org. Chem., 1993, 58, 3731–3735. 152. Shafiee, A., Motamedi, H. and King, A., Purification, characterization and immobilization of an NADPH-dependent enzyme involved in the chiral specific reduction of the keto ester M, an intermediate in the synthesis of an anti-asthma drug, montelukast, from Microbacterium campoquemadoensis (MB5614). Appl. Microbiol. Biotechnol., 1998, 49, 709–717. 153. Ulijn, R,V., De Martin, L., Gardossi L. and Halling, P.J., Biocatalysis in reaction mixtures with undissolved solid substrates and products. Curr. Org. Chem., 2003, 7, 1333–1346. 154. Rozzell, D.,Enzymatic production of the key montelukast intermediate. Spec. Chem.Mag., 2008, April, 36–38. 155. Reddy, G.K., Gralla, R.J. and Hesketh, P.J., Novel neurokinin-1 antagonists as antiemetics for the treatment of chemotherapy-induced emesis. Support. Cancer Ther., 2006, 3, 140–142. 156. Gelo-Pujic, M., Le Guyader, F. and Schlama, T., Microbial and homogenous asymmetric catalysis in the reduction of 1-[3,5-bis(trifluoromethyl)phenyl]ethanone. Tetrahedron Asymm. 2006, 17, 2000–2005. 157. Pollard, D., Truppo, M., Pollard, J., Chen, C.-Y. and Moore, J., Effective synthesis of (S)-3,5bistrifluoromethylphenyl ethanol by asymmetric enzymatic reduction. Tetrahedron Asymm. 2006, 17, 554–559. 158. Van Langen, L.M., van Rantwijk, F. and Sheldon, R.A., Enzymatic hydrocyanation of a sterically hindered aldehyde. Optimization of a chemoenzymatic procedure for (R)-2-chloromandelic acid. Org. Proc. Res. Dev., 2003, 7, 828–831. 159. (a) Gijsen, H.J.M., Wong, C.-H., Unprecedented asymmetric aldol reactions with three aldehyde substrates catalyzed by 2-deoxyribose-5-phosphate aldolase. J. Am. Chem. Soc., 1994, 116, 8422– 8423; (b) Wong, C.-H., Garcia Junceda, E., Chen, L., Blanco, O., Gijsen, H.J.M. and Steensma, D.H., Recombinant 2-deoxyribose-5-phosphate aldolase in organic synthesis: use of sequential two-substrate and three-substrate aldol reactions. J. Am. Chem. Soc., 1995, 117, 3333–3339; (c) DeSantis, G., Liu, J., Clark, D.P., Heine, A., Wilson, I.A. and Wong, C.-H., Structure-based mutagenesis approaches toward expanding the substrate specificity of D-2-deoxyribose-5phosphate aldolase. Bioorg. Med. Chem., 2003, 11, 43–52; (d) Liu, J., Hsu, C.-C. and Wong, C.-H., Sequential aldol condensation catalyzed by DERA mutant Ser238Asp and a formal total synthesis of atorvastatin. Tetrahedron Lett., 2004, 45, 2439–2441. 160. Hu, S., Tao, J. and Xie, Z., Process for producing atorvastatin, pharmaceutically acceptable salts thereof and intermediates thereof. PTC Int. Appl., 2006, WO 2006/134482 A1. 161. Liu, J. and Wong C.-H., Aldolase-catalyzed asymmetric synthesis of novel pyranose synthons as a new entry to heterocycles and epothilones. Angew. Chem. Int. Ed., 2002, 41, 1404– 1407. 162. Samland, A.K. and Sprenger, G.A., Microbial aldolases as C–C bonding enzymes – unknown treasures and new developments. Appl. Microbiol. Biotechnol., 2006, 71, 253–264.

References

79

163. Schu¨rmann, M. and Sprenger, G.A., Fructose-6-phosphate aldolase is a novel Class I aldolase from Escherichia coli and is related to a novel group of bacterial transaldolases. J. Biol. Chem., 2001, 276, 11055–11061. 164. Sugiyama, M., Hong, Z., Whalen, L.J., Greenberg, W.A. and Wong, C.-H., Borate as a phosphate ester mimic in aldolase-catalyzed reactions: practical synthesis of L-fructose and L-iminocyclitols. Adv. Synth. Catal. 2006, 348, 2555 – 2559. 165. Griebenow, K. and Klibanov, A.M., On protein denaturation in aqueous–organic mixtures but not in pure organic solvents. J. Am. Chem. Soc., 1996, 118, 11695–11699. 166. For some recent reviews on the use of enzymes in nonconventional media, see: (a) Dreyer, S., Lembrecht, J., Schumacher, J. and Kragl, U., Enzyme catalysis in nonaqueous media: past, present, and future in biocatalysis in the pharmaceutical and biotechnology industries, 2007, CRC Press, pp. 791–827; . (b) Torres, S. and Castro, G.R., Non-aqueous biocatalysis in homogeneous solvent systems. Food Technol. Biotechnol., 2004, 42, 271–277; (c) Carrea, G. and Riva, S., Properties and synthetic applications of enzymes in organic solvent. Angew. Chem. Int. Ed., 2000, 39, 2226–2254. 167. Butler, L.G., Enzymes in non-aqueous solvents. Enzyme Microb. Technol., 1979, 1, 253–259. 168. Gerard A. Sellek, G.A. and Chaudhuri, J.B., Biocatalysis in organic media using enzymes from extremophiles. Enzyme Microb. Technol., 1999, 25, 471–482. 169. Van Rantwijk, F. and Sheldon, R.A., Biocatalysis in ionic liquids. Chem. Rev., 2007, 107, 2757–2785. 170. Hobbs, H.R. and Thomas, N.R., Biocatalysis in supercritical fluids, in fluorous solvents, and under solvent-free conditions. Chem. Rev., 2007, 107, 2786–2820. 171. Kazlauskas, R.J., Enhancing catalytic promiscuity for biocatalysis. Curr. Opin. Chem. Biol., 2005, 9, 1–7. 172. Gruber-Khadjawi, M., Purkarthofer, T., Skranc, W. and Griengl, H., hydroxynitrile lyase catalyzed enzymatic nitroaldol (Henry) reaction. Adv. Synth. Catal. 2007, 349, 1445–1450. 173. (a) Torre, O., Alfonso, I. and Gotor, V., Lipase catalysed Michael addition of secondary amines to acrylonitrile. Chem. Commun., 2004, 1724–1725; (b) Cai, Y., Yao, S.-P., Wu, Q. and Lin, X.F., Michael addition of imidazole with acrylates catalyzed by alkaline protease from Bacillus subtilis in organic media. Biotechnol. Lett., 2004, 26, 525–528. 174. Zaks, A. and Klibanov, A.M., Enzymatic catalysis in organic media at 100 °C. Science, 1984, 224, 1249–1251. 175. Griebenow, K. and Barletta, G., Dehydrated protein powders as biocatalysts in nonaqueous solvents. In Lyophilization of Biopharmaceuticals (Biotechnology: Pharmaceutical Aspects), Costantino, H.R. and Pikal, M.J. (eds). AAPS Press: Arlington, VA, 2004, pp. 643–668. 176. (a) Klibanov, A.M., Why are enzymes less active in organic solvents than in water? Trends Biotechnol. 1997, 15, 97–101; (b) Klibanov, A.M., Improving enzymes by using them in organic solvents. Nature, 2001, 409, 241–246. 177. Serdakowski, A.L. and Dordick, J.S., Enzyme activation for organic solvents made easy. Trends Biotechnol. 2008, 26, 48–54. 178. Partridge, J., Dennison, P.R., Moore, B.D. and Halling, P.J., Activity and mobility of subtilisin in low water organic media: hydration is more important than solvent dielectric. Biochim. Biophys. Acta, 1998, 1386, 79–89. 179. Bell, G., Halling, P.J., May, L., Moore, B.D., Robb, D.A., Ulijn, R. and Valivety, R.H., Methods for measurement and control of water in nonaqueous biocatalysis. In Methods in Biotechnology, Vol. 15: Enzymes in Nonaqueous Solvents: Methods and Protocols, Vulfson, E.N., Halling, P.J. and Holland, H.L. (eds). Humana Press: Totowa, NJ, 2001, pp. 105–126. 180. Valivety, R.H., Halling, P.J., Peilow, A.D. and Macrae, A.R., Lipases from different sources vary widely in dependence of catalytic activity on water activity. Biochim. Biophys. Acta, 1992, 1122, 143–146. 181. Halling, P.J., What can we learn by studying enzymes in non-aqueous media? Phil. Trans. R. Soc. Lond. Ser. B, 2004, 359, 1287–1297. 182. Bell, G., Halling, P.J., Moore, B.D., Partridge, J. and Rees, D.G., Biocatalyst behaviour in lowwater systems. Trends Biotechnol., 1995, 13, 468–473.

80


183. (a) Rich, J.O., Mozhaev, V.V., Dordick, J.S., Clark, D.S. and Khmelnitsky, Y.L., Molecular imprinting of enzymes with water-insoluble ligands for nonaqueous biocatalysis. J. Am. Chem. Soc., 2002, 124, 5254–5255 and references cited therein; . (b) Carrea, G., Ottolina, G. and Riva, S., Role of solvents in the control of enzyme selectivity in organic media. Trends Biotechnol. 1995, 13, 63–70 and references cited therein. 184. Zacharis, E., Moore, B.D. and Halling, P.J., Control of enzyme activity in organic media by solid-state acid–base buffers. J. Am. Chem. Soc., 1997, 119, 12396–12397. 185. Zaks, A. and Klibanov, A.M., Enzyme-catalyzed processes in organic solvents. Proc. Natl. Acad. Sci., 1985, 42, 3192–3196. 186. Laane, C., Boeren, S., Vos, K. and Veeger, C., Rules for optimization of biocatalysis in organic solvents. Biotechnol. Bioeng., 1987, 30, 81–87. 187. Halling, P.J., Thermodynamic predictions for biocatalysis in nonconventional media: theory, tests, and recommendations for experimental design and analysis. Enzyme Microb. Technol., 1994, 16, 178–206. 188. Secundo, F., Riva, S. and Carrea, G., Effects of medium and of reaction conditions on the enantioselectivity of lipases in organic solvents and possible rationales, Tetrahedron Asymm. 1992, 3, 267–280. 189. Keeling, S.P., Campbell, I.B., Coe, D.M., Cooper, T.W.J., Hardy, G.W., Jack, T.I., Jones, H.T., Needham, D., Shipley, T.J., Skone, P.A., Sutton, P.W., Weingarten G.A. and Macdonald, S.J.F., Efficient synthesis of an -trifluoromethyl--tosyloxymethyl epoxide enabling stepwise double functionalisation to afford CF3-substituted tertiary alcohols. Tetrahedron Lett. 2008, 49, 5101. 190. Roberge, C., Fleitz, F., Pollard, D. and Devine, P., Asymmetric synthesis of cyanohydrin derived from pyridine aldehyde with cross-linked aggregates of hydroxynitrile lyases. Tetrahedron Lett., 2007, 48, 1473–1477. 191. Hirose, Y., Kariya, K., Sasaki, I., Kurono, Y., Ebiike, H. and Achiwa, K., Drastic solvent effect on lipase-catalysed enantioselective hydroloysis of prochiral 1,4-dihydropyridines. Tetrahedron Lett., 1992, 33, 7157. 192. Ballesteros, A., van Beynum, G., Borud, O. and Buchholz, K., Guidelines for the characterization of immobilized biocatalysts. Enzyme Microb. Technol., 1983, 5, 304–307. ¨ .M., Dunn, I.J. and Heinzle, E., Immobilized 193. For some recent reviews, see: Prˇenosil, J.E., Kut, O biocatalysts. In Ullman’s Biotechnology and Biochemical Engineering, vol. 2. Wiley–VCH, Weinheim, 2007, pp. 683–734; Sheldon, R.A., Enzyme immobilization: the quest for optimum performance. Adv. Synth. Catal., 2007, 349, 1289–1307; End, N. and Scho¨ning, K.-U., Immobilized biocatalysts in industrial research and production. Topics Curr. Chem., 2004, 242, 273–317; Bornscheuer, U.T., Immobilizing enzymes: how to create more suitable biocatalysts. Angew. Chem. Int. Ed., 2003, 42, 3336–3337; Cao, L. Immobilised enzymes: science or art? Curr. Opin. Chem. Biol., 2005, 9, 217–226. 194. Schmid, R.D. and Verger, R., Lipases: interfacial enzymes with attractive applications. Angew. Chem. Int. Ed., 1998, 37, 1608–1633; Hasan, F., Shah, A.A. and Hameed, A., Industrial applications of microbial lipases. Enzyme. Microb. Technol., 2006, 39, 235–251. 195. Gonza´lez-Navarro, H., Carmen Banõ´, M. and Abad, C., The closed/open model for lipase activation. Addressing intermediate active forms of fungal enzymes by trapping of conformers in water-restricted environments. Biochemistry, 2001, 40, 3174–3183. 196. Katchalski-Katzir, E. and Kraemer, D.M., EupergitÒ C, a carrier for immobilization of enzymes of industrial potential. J. Mol. Catal. B, 2000, 10, 157–176. 197. Roy, J.J. and Abraham, T.E., Strategies in making cross-linked enzyme crystals. Chem. Rev., 2004, 104, 3705–3721. 198. Weetall, H.H. and Pitcher Jr, W.H., Scaling up an immobilized enzyme system. Science, 1986, 232, 1396–1403. 199. Schoevaart, R., Wolbers, M.W., Golubovic, M., Ottens, M., Kieboom, A.P.G., van Rantwijk, F., van der Wielen, L.A.M. and Sheldon, R.A., Preparation, optimization, and structures of cross-linked enzyme aggregates (CLEAs). Biotechnol. Bioeng., 2004, 87, 754–762. 200. Mateo, C., Palomo, J.M., van Lancen, L.M., van Rantwijk, F. and Sheldon, R.A., A new, mild cross-linking methodology to prepare cross-linked enzyme aggregates. Biotech. Bioeng., 2004, 86, 273–276.

References

81

201. Dunker, A.K. and Fernańdez, A., Engineering productive enzyme confinement. Trends Biotechnol., 2007, 25, 189–190. 202. Pierre, A.C., The sol–gel encapsulation of enzymes. Biocatal. Biotrans., 2004, 22, 145–170 and references cited therein. 203. Reetz, M.T., Tielmann, P., Wiesenho¨fer, W., Ko¨nen, W. and Zonta, A., Second generation sol– gel encapsulated lipases: robust heterogeneous biocatalysts. Adv. Synth. Catal., 2003, 345, 717–728. 204. Lu¨tz, S., Rao, N.N. and Wandrey, C., Membranes in biotechnology. Chem. Eng. Technol., 2006, 29, 1404–1415; Gekas, V.C., Artificial membranes as carriers for the immobilization of biocatalysts. Enzyme Microb. Technol., 1986, 8, 450–460. 205. Anastas, P.T. and Warner, J.C., Green Chemistry: Theory and Practice, Oxford University Press, 2000. 206. Sheldon, R.A., Organic synthesis - past, present and future. Chem. Ind., 1992, 23, 903–906. 207. Constable, D.J.C., Curzons, A.D. and Cunningham, V.L., Metrics to ‘green chemistry’ – which are the best? Green Chem. 2002, 4, 521–527. 208. ACS GCI Pharmaceutical Roundtable, http://portal.acs.org/portal/acs/corg/content?_ nfpb¼true&_pageLabel¼PP_TRANSITIONMAIN&node_id¼1422&use_sec¼false&sec_ url_var¼region1, (last access 24 June 2008). 209. Constable, D.J.C., Curzons, A.D., Freitas dos Santos, L.M., Geen, G.R., Kitteringham, J., Smith, P., Hannah, R.E., McGuire, M.A., Webb, R.L., Yu, M., Hayler, J.D. and Richardson, J.E., Green chemistry measures for process research and development. Green Chem., 2001, 3, 7–9; Curzons, A.D., Constable, D.J.C., Mortimera, D.N. and Cunningham, V.L., So you think your process is green, how do you know? Using principles of sustainability to determine what is green – a corporate perspective. Green Chem., 2001, 3, 1–6. 210. Jimeńez Gonza´lez, C., Curzons, A.D., Constable, D.J.C. and Cunningham, V.L., Cradle-to-gate life cycle inventory and assessment of pharmaceutical compounds. Int. J. Life Cycle Assess., 2004, 9, 115–121; Curzons, A.D., Jimeńez Gonza´lez, C., Duncan, A.L., Constable, D.J.C. and Cunningham, V.L., Fast life cycle assessment of synthetic chemistry (FLASC) tool. Int. J. Life Cycle Assess., 2007, 12, 272–280; Jimeńez Gonza´lez, C., Overcash, M.R. and Curzons, A., Waste treatment modules – a partial life cycle inventory. J. Chem. Technol. Biotechnol., 2001, 76, 707–716. 211. Jimeńez Gonza´lez, C., Constable, D.J.C., Curzons, A.D. and Cunningham, V.L., Developing GSK’s green technology guidance: methodology for case-scenario comparison of technologies. Clean Technol. Environ. Pollut., 2002, 4, 44–53. 212. Sheldon, R.A., The E factor: fifteen years on. Green Chem., 2007, 9, 1273–1283. 213. Menzel, A., Werner, H., Altenbuchner, J. and Groeger, H., From enzymes to ‘designer bugs’ in reductive amination: a new process for the synthesis of L-tert-leucine using a whole cellcatalyst. Eng. Life Sci., 2004, 4, 573–576; Bruggink, A., Schoevaart, R. and Kieboom, T., Concepts of nature in organic synthesis: cascade catalysis and multistep conversions in concert. Org. Proc. Res. Dev., 2003, 7, 622–640; Buckland, B.C., Robinson, D.K. and Chartrain, M., Biocatalysis for pharmaceuticals – status and prospects for a key technology. Metab. Eng., 2000, 2, 42–48. 214. Bacher, A., Stohler, P. and Weber, W., Process for the production of lipstatin and tetrahydrolipstatin. Eur. Pat. Appl., 2002, EP803576 A2. 215. (a) Fleming, M.P., Han, Y.-K., Hodges, L.M., Johnston, D.A., Micheli, R.P., Puentener, K., Roberts, C.R., Scalone, M., Schwindt, M.A. and Topping, R.J., Process for the (enantioselective) synthesis of 3,6-dialkyl-5,6-dihydro-4-hydroxy-pyran-2-ones via intramolecular cyclization of (homochiral) -halo esters. PCT Int. Appl., 2001, WO 2001057014 A2; (b) Birk, R., Karpf, M., Pu¨ntener, K., Scalone, M., Schwindt, M. and Zutter, U., With asymmetric hydrogenation towards a new, enantioselective synthesis of orlistat. Chimia, 2006, 60, 561– 565; (c) Schwindt, M.A., Fleming, M.P., Han, Y.-K., Hodges, L.M., Johnston, D.A., Micheli, R.P., Roberts, C.R., Snyder, R., Topping, R.J., Pu¨ntener, K. and Scalone, M., enantioselective synthesis of a key intermediate in a new process for orlistat using asymmetric hydrogenation and a Grignard reagent promoted lactone cyclization. Org. Proc. Res. Dev., 2007, 11, 524–533.

82


216. Karpf, M. and Zutter, U., Process for the preparation of oxetanones. Eur. Pat. Appl., 1991, EP 443449 A2. 217. The chemical route to orlistat was ultimately favoured over the biosynthetic route as the latter required complex downstream processing that eliminated the benefits gained from the biotransformation itself. More recent advances in metabolic engineering and drownstream processing might have resulted in the development of a more competitive process. 218. Poechlauer, P. and Wagner, M., Enzymatic process to separate racemic mixtures of delta valerolactones. US PCT Appl., 1995, US 5412110. 219. Scott, A.I., Discovering nature’s diverse pathways to vitamin B12: a 35-year odyssey. J. Org. Chem., 2003, 68, 2529–2539. 220. Bornscheuer, U.T. and Buchholz, K., Highlights in biocatalysis – historical landmarks and current trends. Eng. Life Sci., 2005, 5, 309.

2 Biocatalyst Identification and Scale-up: Molecular Biology for Chemists Kathleen H. McClean

2.1

History of Biotechnology

Biotechnology is a modern discipline with ancient origins. Fermentation has been exploited by humans for more than 6000 years for the manufacture and preservation of foodstuffs. The modern-day manufacture of beer, bread, wine, dairy products, and fermented bean products are the descendants of these early experiments in food processing. Over the past 200–300 years many of these processes have made the transition from traditional or cottage industries to large (industrial)-scale, controlled manufacturing processes. This was facilitated by the development of the discipline of microbiology, which brought a better understanding of the identity and behaviour of the microorganisms involved. Several of the bacteria, yeasts and filamentous fungi associated with these traditional processes are still work-horses of modern biotechnology – organisms such as Saccharomyces cerevisiae, lactic acid bacteria and Aspergillus oryzae. Many have been used in industrial fermentations to produce a diverse range of products, such as vitamins, amino acids, organic acids and solvents (Figure. 2.1). ‘White biotechnology’ had begun to emerge as a means to produce platform chemicals, biofuels and chemical intermediates. Early examples of biotransformation using microbes and defined chemical substrates began to become established in the mid-nineteenth century. Pasteur noted in 1858 that when a solution of an ammonium salt of (–)-tartaric acid was fed to a culture of the mould Penicillium glaucum the (þ)-tartaric acid was consumed, leaving the ()-tartaric acid,

Practical Methods for Biocatalysis and Biotransformations 2009 John Wiley & Sons, Ltd


84

Biocatalyst Identification and Scale-up: Molecular Biology for Chemists SOLVENTS AND BIOFUELS Ethanol CH3CH2OH Butanol CH3CH2CH2CH2OH

A ceti c A ci d CH3CO2H 1,3-Propanediol HOCH2CH2CH2OH

FINE CHEMICALS, POLYMERS AND PHARMACEUTICAL INTERMEDIATES Me

CO2H

HO

CO2H

(E) (R)

CO2H

O HO

O n

Itaconic Acid

(S) (R)

NH2 Aminoshikimic Acid

Poly-Lactic Acid

VITAMINS

O

OH Me HO

N

(E)

NH

O

O Me

H

N

(E)

N OH O

(R) (R)

HO OH Vitamin C ANTIBIOTICS Me

NMe2

OH (S)

(S) (Z)

OH O Tetracycline

OH

(S)

Riboflavin Vitamin B2 OH

(Z)

(S)

OH

OH

CONH2

H N

Ph O

OH

S N

OH

Me Me

O CO2H

O Penicillin G

Figure 2.1 Structures of various industrial fermentation products

thus providing a very early example of a whole-cell biotransformation. The pioneering work of Eduard Buchner (1897) demonstrated the fermentation of sugar by cell-free extracts of yeast, establishing the principle that fermentation occurred as a result of soluble microbial catalysts (enzymes). This established the important principle that not all biological transformations required living cells. One of the first major pharmaceutical biotransformations was the development of the synthesis of hydrocortisone in the late 1940s by whole-cell hydroxylation1 (Figure 2.2). Up until then a 40-step synthetic route developed by the Noble Prize winning chemist R.B.Woodward was the only source of this important drug substance and intermediate.2 Nowadays, a biocatalyst exists for the selective hydroxylation of every position on the steroid nucleus.3 Biotransformations using native organisms (whole-cell or cell extracts) can be limited by poor expression of the specific enzyme, by the presence of more than one enzyme activities competing for the substrate or by the presence of isozymes displaying a range of specificities. However, by 30–40 years ago, significant advances in microbial genetics had paved the way for recombinant DNA (rDNA) technology which was to result in radical and

2.2 Identifying Potential Biocatalysts for Chemical Synthesis O

O Me Me

O

85

Me

OH OH

HO

Whole Cells

OH OH

Me

O

Figure 2.2 Regio- and stereo-selective steroid hydroxylation

rapid advances in biotechnology. New techniques for manipulating DNA emerged, such as the development of vectors and cloning strategies, which allowed rapid generation of novel recombinant strains of microbes which could produce new enzymes. Many of the first commercial products generated using these technologies were mass-produced extracellular hydrolase enzymes (such as proteases: subtilisin, thermolysin and chymotrypsin). Often, these ‘bulk enzymes’ were originally developed for consumer products or process industry applications and were also exploited by chemists in synthetic reactions and for the resolution of racemates.4 Rapid methods of random and specific site-directed mutagenesis coupled with screening became routine in the enzyme manufacturing sector, enlarging the portfolio of ‘process’ enzymes which could also be exploited for their biocatalytic (synthetic) potential. The development of rDNA technology also accelerated research in the regulation of genetic processes, and the development of technologies such as polymerase chain reaction (PCR) and automated gene sequencing have in turn helped to drive advances in our understanding of how genetic information relates to function (bioinformatics). As a result, the tremendous rate of expansion of information and the accessibility of rDNA technologies (including robotics and advanced bioinformatics) has meant that screening and genetic modification procedures which a decade ago could have taken months or years can now often be completed in weeks or even days. This led to the increased production of enzymes specifically for biotransformation applications. The number of novel enzymes or enzyme genes identified has risen exponentially in the past 10 years, and a greater number and diversity of biocatalysts are available or accessible for synthetic chemistry (in the same decade, a 35-fold increase in the number of articles with ‘biocatalysis’ in the title or abstract has been recorded by bibliographic resources such as PubMed).

2.2

Identifying Potential Biocatalysts for Chemical Synthesis

Biological systems can offer many attractive features as catalysts for the synthetic chemist, such as high substrate specificity, precise stereo- and regio-selectivity and mild reaction conditions. The diversity of biochemical reactions is an indicator of the potential of enzymes in synthetic applications. Therefore, when designing a synthetic route to a target molecule, a number of steps might best be performed using a biocatalyst, particularly where the generation of chirality is involved, and so biocatalysis should be routinely considered. This section briefly outlines the options available to the researcher when contemplating using a biocatalyst.

86

2.2.1

Biocatalyst Identification and Scale-up: Molecular Biology for Chemists

Literature Precedents

As with chemical synthesis, the first step when prospecting for a particular biotransformation is to perform a literature search to check whether a suitable precedent has been described. Extensive technical literature resources in the public domain provide both examples of specific enzyme-catalysed reactions and descriptions of transformations where enzyme activity is inferred if not explicitly described. Currently, searches of online databases such as PubMed reveal over 2000 new publications per annum in the subject of enzyme catalysis (excluding reviews). In some fortunate instances, an exact match for the desired biotransformation might be available in the literature that uses a commercial enzyme or microbial strain that is accessible through culture collections and provides sufficient activity for use in preparative-scale reactions. 2.2.2

Commercial Enzyme Sources

When the reaction of interest is catalysed by a stable enzyme and a single biotransformation step is required, it is often possible to find a suitable commercial enzyme for this step. Some commonly used enzymes can be sourced from general supply houses such as Sigma– Aldrich, but there are now several specialist suppliers who provide enzymes for biocatalysis applications. At the time of writing, the most comprehensive catalogue of enzymes is probably that supplied by Codexis (www.codexis.com). Other major suppliers include Meito Sangyo (www5.mediagalaxy.co.jp/meito/), Amano Enzyme (www.amano-enzyme. co.jp/eng/company), ChiralVision (www.chiralvision.com) and Enzysource (www. enzysource.com ). There are many other smaller suppliers, some of which also offer a more ‘bespoke’ service providing a particular class of enzyme, or supplying additional services such as enzyme formulation, immobilization or enzyme kits. Several academic groups also maintain up-to-date pages of links to enzyme suppliers; these include the CoEBio3 site at the University of Manchester (www.coebio3.org) and the biocatalysis group pages at the University of Graz (http://borgc185.kfunigraz.ac.at/). Many enzymes require the presence of small organic non-protein groups (cofactors) in order to catalyse reactions. Dependence on cofactors may limit the usefulness of the enzyme if the cofactors are expensive, unstable or difficult to recycle. This becomes an issue when using redox enzymes such as dehydrogenases, which often require the presence of the reduced form of nicotinamide adenine dinucleotide (NADH) or its phosphorylated analogue (NADPH). Enzyme and cofactor recycling itself can be avoided by using wholecell systems, or the cofactor can be recycled in vitro by the action of a second enzyme and the inclusion of a suitable substrate which is oxidized. An example is the use of formate dehydrogenase (commercially available) for the oxidation of formic acid to CO2 for the recycling of NADH from NADþ. It should be noted that some commercial enzyme preparations may contain several enzyme isomers (enzymes derived from one source which belong to the same enzyme class but differ in specificity, stability or other properties). This is most often the case when the commercial preparation was developed for a process industry application rather than a specific chemical biotransformation application. Some fungal enzymes, such as laccase, are sometimes supplied as crude enzyme mixtures. Fungal laccases are manufactured on a huge scale (multitonne per annum) and are principally used in bulk processes such as wood

2.2 Identifying Potential Biocatalysts for Chemical Synthesis

87

pulp processing and textile dye bleaching. If a very specific activity is required then the chemist may need to request additional information from the supplier or manufacturer, or be prepared to screen several commercial preparations. However, the growing trend to produce enzymes using rDNA technology is reducing the frequency of these problems. 2.2.3

Culture Collections as Sources of Microorganisms

If the precise original biological source of the desired enzyme for a precedented transformation is known (say, from a microbe with a stock centre code or number), then it can be easy to obtain that microbe and grow it according to the defined conditions in the literature reference or stock centre recommendations. There are more than 500 registered microbial stock centres worldwide where microbial cultures and cell lines are curated. Most of these are accessible to the public, and samples of cultures or cell lines can be obtained for relatively modest fees. Further information on stock centres, including contact details, is available from the World Federation for Culture Collections (www.wfcc.info), which provides links to culture collections. Although most developed countries have national (and sometimes specialized) collections, there are a few major collections which are most frequently used for the deposit of microbes of industrial interest (Table 2.1). These collections are a vital resource for biotechnology, as they provide access to certified pure microbial cultures as well as being valuable technical resources for microbial identification with information on culture and characteristics of the microbes. In addition, they provide safe repositories for materials covered by patent protection. Alongside the large and often diverse culture collections there are many specialized collections which

Table 2.1 Some major microbial culture collections. Most of the larger collections have online searchable catalogues and provide other important information on pathogenicity, cell culture and maintenance, as well as bibliographic information relating to individual strains Culture collection

Abbreviation

Web address

Major collections

Belgian Coordinated Collections of Microorganisms

BCCM

www.bccm.belspo.be

Bacteria, Fungi Yeasts DNA

Deutsche Sammlung von Mikroorganismen und Zellkulturen

DSMZ

www.dsmz.de

Bacteria, fungi, yeasts, cell lines, DNA, viruses, Archaea

American Type Culture Collection

ATCC

www.atcc.org

Bacteria, fungi, yeasts, cell lines, DNA, viruses, Archaea

Centraalbureau voor Schimmelcultures

CBS

www.cbs.knaw.nl

National Collection of Industrial Bacteria

NCIMB

www.ncimb.com

Fungi, yeasts, CBS also host NCCB (Netherlands Bacterial Culture Collection) Bacteria (industrial food, environmental and marine), DNA resources

88


provide resources (materials and information) associated with one type or strain of microbe (such as the Escherichia coli genetic stock centre: www.cgsc.biology.yale.edu). Many of the larger commercial collections also have excellent online catalogues which can also be searched using relevant keywords for recorded enzyme activity, metabolic pathways, substrates or products, or their environmental origin. As the interest in using culture collections as biocatalyst resources has increased, so the annotation of the catalogues has improved, and most larger collections actively seek and collate information documenting their strain’s biocatalytic capabilities. What can be done if there is not an exact literature precedent for the required enzymatic transformation? In these cases some sort of screening procedure may be necessary, either by direct screening of commercial enzymes, culture collections and environmental isolates, or by using a bioinformatics-based approach to identify potential enzymes based on sequence information. 2.2.4

Enzyme and Gene Databases, Bioinformatics and the Search for New Enzymes

Searching for information on enzymes can be made a lot easier by surveying the collated information available at online databases dedicated to genetic information (sequence based) and to enzymes (usually based on analysis of observed activity). The genetic (sequence) information can refer to the gene sequences of cloned enzymes where activity has been demonstrated, and also to the predicted enzyme genes ‘mined’ from the huge resource of genetic information accrued from genome sequencing projects and metabolomics experiments (the direct recovery of DNA from the environment). Originally, information on genetic resources and enzyme activity was often collected independently, but nowadays these resources are often integrated or cross-referenced. The correlation between information relating to proteins (enzymes) and nucleic acid sequences illustrates the increasing importance of bioinformatics – the application of mathematical and computing techniques to interpretation of sequence information. A good practical starting point is to look at databases which primarily deal with demonstrated enzyme activity, since this may identify enzymes which could be relatively easy to obtain. Most of these databases have their origins in sectors of biological research, although increasingly the role of enzymes in biocatalysis is referenced in an accessible form. Unfortunately, not all known enzymes are referenced in every database, and the occurrence in a database may not mean the enzyme has been purified or that it comes from an accessible microbial source. However, by knowing that the databases differ in their focus, this can be used to collect complementary sets of information about individual enzymes. BRENDA (www.brenda-enzymes.org) offers a comprehensive database of enzymes in the academic literature. Much of the information presented focuses on functional information, so that information on substrates, stability, pH range, etc. is particularly easy to obtain. The database covers enzymes from all types of organisms. Searches can be made by many methods: by enzyme name, by class, substrate, product, molecule structure (either by name or by using a graphical interface), and by organism. It is also possible to search amino acid sequence databases for families of sequence-related enzymes, or for sequence motifs, or even to check if a gene is likely to encode an enzyme.5 The University of Minnesota Biocatalysis/Biodegradation Database (http://umbbd.msi.umn.edu/index.html) provides a


89

comprehensive overview of microbial biocatalytic reactions and biodegradation pathways.6 This can be particularly useful if the substrate is a known xenobiotic. In many instances the reactions described by this resource are referenced to original publications and named microbial isolates. It is important to remember that the content of this database focuses on the enzymatic activities of microbes in the environment and the mechanisms they use to degrade organic compounds, including pollutants. The database can be searched by several methods, including by pathway, by chemical compound (including graphical structure search), by organism and by enzyme name. This site has organized biocatalytic reactions into pathways, noting that in natural environments the steps of the biodegradative pathway could exist in a single organism or in a range of microbes (a microbial consortium). Features of this website also include a pathway prediction tool which attempts to generate plausible biodegradation (biotransformation) routes for a given organic compound based on ‘rules’ generated by extensive review of the academic literature. The information is of particular use if one needs to check whether a particular transformation is likely to occur (and may identify an actual candidate enzyme); it is also important when carrying out whole-cell biotransformations, as it can be used to predict the likelihood of unwanted side reactions or downstream modifications of the product. The site also hosts an excellent page of links to other online resources in microbial biotechnology. There are several other online searchable sites which can be used to search for enzymes, and the most significant of these are linked to or embedded in bioinformatics resources. These types of resource are probably more useful when considering obtaining a new enzyme by cloning experiments or when planning mutagenesis experiments to generate enzymes with altered properties. At its most basic, bioinformatics involves using basic sequence analysis tools in the identification of features in nucleic acids or amino acid sequences. Widely accessible programs are routinely used to identify gene coding sequences, features such as regulatory sequences and introns (interrupting noncoding DNA sequences which do not encode polypeptide) and prediction of proteolysis sites in polypeptide sequences. However, the accumulation of large quantities of sequence information, combined with advances in computational methods, has made it possible to perform many more complex analyses. The principle activities in bioinformatics include mapping and analysing DNA and protein sequences, aligning different DNA and protein sequences to compare them (identifying homology) and creating and viewing three-dimensional models of protein structures. All these activities can be applied in the search for new or improved enzymes. One basic approach is to search databases of sequence information for predicted enzyme sequences (sequence search service). Usually, when a DNA or RNA sequence is lodged in a searchable database, the depositor will have performed a basic annotation which may include predictions of enzyme amino acid sequences. Individual gene sequences can be compared with all the other sequences in a database or library in searches for overall sequence homology (e.g. ‘The Basic Local Alignment Search Tool’ (BLAST) finds regions of local similarity between sequences7). These types of analysis can be used to infer functional or evolutionary relationships between sequences. Protein sequence features have been systematically analysed to help identify the specific features or motifs characteristic of protein function (including enzyme activity) using facilities such as PROSITE, a database of entries describing the domains, families and functional sites of proteins, as well as their associated amino acid patterns, signatures and profiles.8 Further information on these and other methods of ‘data mining’ and analyses are given at the

90


ExPASy (Expert Protein Analysis System, www.expasy.ch) or the National Centre for Biotechnology Information (NCBI, www.ncbi.nlm.gov) websites. Both websites provide bioinformatics tools, links to sequence databases and extensive bibliographic resources. As an example of the wealth of information available on individual enzymes, at the time of writing a search based on ‘nitrilase’ in the ‘Entrez protein’ section of NCBI will recover more than 10 000 references to nitrilase enzyme amino acid sequences. These can be rapidly screened online by organism, and the individual entries will have links to amino acid and gene sequence, relevant literature and information on protein features (such as conserved domains). 2.2.5

Metagenomics: Sampling DNA Directly from the Environment

Another approach now available when searching for new enzymes is to obtain the gene for the enzyme from libraries of DNA recovered directly from the environment. This avoids the need to know much about the original microorganism and also eliminates the need to grow it in the laboratory. Much of the interest in metagenomics comes from the discovery that the vast majority of microorganisms had gone unnoticed until relatively recently.9 Traditional microbiological methods rely upon laboratory cultivation of organisms. For quite some time before the development of molecular biology techniques it had already been recognized that many microbes eluded description or characterization because they could not be cultured by standard microbiological techniques. Surveys of ribosomal RNA genes taken directly from the environment revealed that cultivation-based methods find less than 1% of the bacterial and archaeal species in a sample (Archaea are a group of organisms which often resemble bacteria in morphology but have some features which distinguish them from both prokaryotes an eukaryotes). This illustrated the extent of our ignorance about the range of metabolic and species diversity in the microbial world. In the recent past there has been a trend to sample microbes and microbial DNA from ‘mega diversity ecosystems’ typically found in locations such as Mexico, Central America or South East Asia, or extreme environments found in regions of volcanic activity (Hawaii, Iceland), deep ocean thermal vents and permafrost. Sampling DNA from multiple sites probably increases the species diversity represented in the pooled samples. Samples from particular environments might help to recover DNA enriched in genes for enzymes with certain properties – thermostable enzymes could be expected to be found associated with high-temperature environments; carbohydratases might be produced in the digestive tracts of herbivores.10 Similarly, polluted or contaminated sites might be useful locations to look for the DNA-encoding enzymes which act on the contaminants or related substances. For several commercial organizations specializing in the development of novel enzymes for industrial processes, these metagenomic pools are a valuable resource to be ‘mined’ using high-throughput technologies to discover new enzymes (see Figure 2.3).11 These methods are represented in a rather simplistic manner, as with any set of techniques there are some limitations: the DNA may be extensively damaged; it can be difficult to recover entire intact genes on smaller fragments of DNA; redundancy could be an issue where some species predominate; and specialized DNA cloning techniques may be needed to maintain long fragments of genomic DNA. Recently, an additional screening method has been proposed in which catabolic genes induced by various substrates are identified from metagenomic DNA libraries by using automated cell-sorting screening


91

Genomic DNA extraction

DNA fragments

Digested vector

Ligation

Transformation

Sequence driven analysis

Function driven analysis

Figure 2.3 Metagenomic cloning experiments. Isolation of genomic DNA directly from environments (soil, plants, mixed environments or thermal-vent worms are the examples illustrated here) can recover DNA fragments which could encode for enzymes. The DNA fragments can be ligated to plasmids or DNA linkers, and then subjected to functional screening (expression cloning) and/or sequence analysis. Amplification by PCR can sometimes be used to yield libraries enriched with clones containing selected sequence motifs relating to families of enzymes

techniques (such as fluorescence activated cell sorting).12 This method was applied successfully to isolate aromatic hydrocarbon-induced genes from a metagenomic library. The challenges are to identify a subset of relevant clones from very large libraries of random DNA clones. Nevertheless, metagenomics offers a methodology to sample true enzymatic biodiversity which is several orders of magnitude greater than that which could be found using ‘conventional’ microbiology. In some senses, metagenomic screening is a reflection on, and expansion of, the historic extensive screening of soil samples for culturable microbes which could produce interesting secondary metabolites. The techniques now also sample the unculturable organisms and can search for potential enzymes using sequence-based methodologies as well as by demonstration of activity by expression cloning. There remain the difficulties which are

92


sometimes encountered when trying to express an active enzyme (some of these will be discussed in later sections of this chapter – protein truncation, presence of introns, misfolding, post-translational modifications, codon matching, inappropriate expression host, etc.). The more fruitful approach may be first to recover and analyse the sequences from the environment, identify the promising clones (e.g. those which have sequence homology to known enzymes), then develop strategies for expression cloning (to produce the enzyme in a host cell such as E. coli) and functional testing of the enzymes. This has recently been used by several groups as a method to discover new enzymes involved in polyketide synthesis.13 However, where there is access to existing high-throughput functional screening methods (such as growth on single substrate), the direct functional screen can also be useful.14 High-throughput functional screening systems typically employ colorimetric or fluorescence-based assays to demonstrate activity, and often use robotic systems to generate and analyse the many thousands of tests required in screening experiments. Bacteria-like microbes found in extreme environments (‘extremophiles’) were sometimes presumed to belong exclusively to the specialized ‘domain’ known as Archaea. However, it has transpired that the arrangement of microbes in ecological niches was not as simple as this – many Archaea have been found in temperate zones and in ecosystems similar to those occupied by bacteria, and some ‘conventional’ microbes occupy extreme environments. Nevertheless enzymes isolated directly or indirectly from organisms found in extreme environments are likely to harbour useful adaptations which could be exploited in industrial processes (such as solvent tolerance and thermostability). As an example, the readily available and versatile lipase B from Candida antarctica (CAL-B) enzyme, isolated from an extremely cold environment (a lake on the Antarctic continent), surprisingly shows remarkable thermal tolerance under certain conditions, notably in organic solvents, where it can often be used at 60 C. Structural studies of these enzymes can yield insights on the features which confer these advantages, information which might be used in the future to inform targeted evolution or modification of other enzymes, as well as contributing to the general pool of known enzymes.15 Novel enzymes obtained via metagenomics may still need to undergo performance optimization by molecular biology or protein engineering to make them suited for industrial processes.

2.3

Molecular Biology for Improved Biocatalysts

Many reported biotransformations are initially only demonstrated on a very small scale, the substrates or products may be subject to competing reactions if other enzymes are present (this can be a serious issue in whole-cell biocatalysis), or the desired enzyme is insufficiently active or produced in low levels. For many biotransformations a little care and attention is needed in the growth of the microbe to achieve the desired results. Production of a specific enzyme from a microbe can often be increased by growing the cells in the presence of a very small concentration (typically micromolar) of an inducer. The inducer could be a ‘natural’ enzyme substrate, a substrate mimic or a molecule which is in some way associated with a substrate’s availability or role in metabolism. This process is called induction and represents a genetic ‘switch’ which cells use to respond

2.3 Molecular Biology for Improved Biocatalysts

93

to environmental changes, and can be used to control enzyme production both by wild-type cells and when applied in rDNA technology. A typical example would be the induction of lipase production by the presence of fats,16 whilst providing a protein substrate or starving the cell of nitrogen could be a method to stimulate protease production.17 In contrast, enzymes which are involved in essential tasks, such as central metabolic functions, are available all the time to the cell – their genes are often referred to as housekeeping genes and their constant level of enzyme production is known as constitutive expression. These systems controlling levels of enzyme production are exploited in rDNA technology. Controllable (inducible) enzyme production systems are generally used to produce maximal yields of functional biocatalyst. For laboratory-scale production of enzymes using rDNA technology in E. coli, many commonly used expression systems use isopropyl- -D-thiogalactopyranoside (IPTG) as an inducer. Although convenient for small-scale production, IPTG is rather expensive and toxic, often making it unsuitable for industrial-scale manufacture of enzymes. Luckily, several alternative inducible control systems are available. A typical example of an alternative system is the use of the relatively cheap and nontoxic sugar arabinose as an inducer of gene expression in E. coli.18 Another strategy is to link the induction of enzyme production to predictable changes in cell physiology, such as the change from the exponential growth phase to the stationary phase. This has been demonstrated in industrial strains of Streptomyces lividans, where the gene of interest has been linked to a promoter (a genetic regulatory unit) which is only active in the stationary phase.19 Difficulties sometimes arise if the enzyme of interest is derived from an organism which is difficult or impossible to grow using conventional laboratory facilities – organisms such as extremophiles (requiring extreme temperatures, pressure or salt levels for culture), or fastidious organisms which have complex (expensive) growth requirements, or from a multicellular organisms which cannot readily be obtained or maintained in a conventional microbiology laboratory. Where the enzyme is from a mammalian source there can be additional safety, ethical or regulatory problems if the biotransformation is destined for pharmaceutical manufacture (since the catalyst will probably be derived from human tissue or slaughterhouse waste), or if consistent quality criteria of the supplies are required which are derived from unregulated animal by-products. An interesting example of this type of material is pig liver esterase (PLE), a versatile biocatalyst which fell into disuse at least in part due to concerns over the safety of animal-derived products. Recently, recombinant PLE enzymes (including commercially produced enzyme) have become available, making its broader use in industrial applications possible once more.20 Enzymes from other plant or animal sources may also be difficult to obtain in quantity, either due to limited access to the source material and/or difficulty in obtaining enough purified active enzyme to perform the reaction. In other cases, a less-than-suitable enzyme is available and one would like to change its properties such as its substrate selectivity or process stability, or it might be thought necessary to find a new enzyme – either by searching for one from nature and/or by engineering of existing enzymes. It is in these instances that rDNA technology becomes an essential tool for producing quantities of usable enzymes manufactured in a controlled manner by a microbial host. This technology aims to extract the DNA from a gene pool and tries to isolate and clone the desired gene(s) for a particular application. For example, the bacterium Rhodococcus erythropolis NCIMB 11540 (from the collection of National Collections of Industrial Food and Marine

94


Bacteria, Aberdeen, UK), was found to have a highly active nitrile hydratase/amidase enzyme system, based on whole-cell biotransformation experiments.21 Subsequently, individual enzymes (nitrile hydratase and amidase) from this strain were cloned and expressed separately in E. coli.22 However, distribution of some strains or other materials from these public collections may be limited, usually as a result of the restrictions on their commercial use imposed by intellectual property rights. How does an understanding of the principles of molecular biology assist in (re)design of biocatalysts? Once the gene specifying an enzyme has been sequenced, the sequence information can be used by the molecular biologist to ‘tailor’ the enzyme using a combination of synthetic and genetic techniques at the DNA sequence level so as to modify the enzyme’s catalytic activity, improve its process compatibility and in some instances improve the actual yield of enzyme manufactured in the fermentation process. It is also important to note that molecular biology, while it is a very powerful tool, is probably most effective in industrial process development when used in conjunction with other techniques such as enzyme formulation, immobilization and appropriate process design engineering.23 rDNA technology has many applications in speciality chemical and pharmaceutical manufacturing beyond the current topic of biocatalysis for small-molecule manufacturing. More advanced related topics include metabolic engineering, advanced fermentation processes and production of biopharmaceuticals, to name but a few. This brief overview of the field will concentrate on the basic principles of rDNA technology for enzyme production, new enzyme discovery and enzyme modification. 2.3.1

Introduction to rDNA Technology (Applied Molecular Biology)

As is the case for many scientific disciplines, molecular biology has developed its own terminology which can appear complex and sometimes bewildering to other scientists. Many acronyms have been developed which help to produce a ‘shorthand’ notation for describing the manipulation of large biomolecules with correspondingly large formal names. For those not familiar with these terminologies, a brief primer of some of the basic principles may be useful. More comprehensive descriptions of the principles of molecular biology are available in several textbooks; for example, see Ref. 24. 2.3.1.1

The Central Dogma of Molecular Biology

The ‘central dogma’ relates to the transfer of sequence-based information in biological systems. DNA – usually the primary store of genetic information and maintained in the cell as double-stranded DNA (dsDNA) molecules – can be faithfully copied (replicated) as new DNA molecules (DNA replication) by an enzyme called DNA-dependent DNA polymerase (usually simply called ‘DNA polymerase’). The sequence-based DNA information can also be copied into messenger RNA (mRNA) by a process called transcription and the resulting mRNA molecules are often referred to as transcripts. Proteins can then be synthesized using the information in mRNA as a template (translation). When inheritable information from a gene, such as the DNA sequence, is made into a functional gene product (such as protein or RNA) the process is known as gene expression and an expression system is required to express it. The basic principles of the central dogma are illustrated in Figure 2.4.


95

DNA Replication

Reverse Transcription

Transcription RNA Translation

Protein

Used by all cells

Used by retroviruses

Figure 2.4 The central dogma of molecular biology – transcription of DNA to RNA to protein. This concept (often called the ‘dogma’) forms the backbone of molecular biology and is represented by four major stages. (1) The DNA replicates its information by conservative replication (by means of DNA polymerases). (2) The DNA codes for the production of mRNA during transcription. In some viruses the primary repository of genetic information is RNA, and the equivalent DNA molecules can be generated by a process known as reverse transcription. (3) In eukaryotic cells, an additional step occurs: the mRNA is processed (essentially by splicing off noncoding regions called introns) and the mature mRNA migrates from the nucleus to the cytoplasm. (4) mRNA carries coded information to specialized complexes of protein and ribonucleic acids called ribosomes. The ribosomes ‘read’ this information and use it for protein synthesis. This process is called translation

The objective of gene cloning is frequently to express protein products from the cloned gene sequences often in a foreign host (i.e. in a cell from a different species). Some special instances of reverse transcription occur in certain viruses, where DNA can be synthesized using RNA as the primary template. The reverse transcriptase system is important when trying to clone and express proteins from eukaryotes (yeasts, fungi, plants and animals), as the organization of their genomic DNA has some important differences from that of prokaryotes (Bacteria and Archaea – see Section 2.3.1.5). Genes from these organisms are frequently cloned as functional units in bacteria by means of in vitro manipulation of the mRNA to recover a DNA sequence which can be used to express proteins in bacteria. 2.3.1.2

Enzyme Tools in Molecular Biology

Several of the enzymes involved in the processes of replicating, transcription and reverse transcription are available commercially and are used by molecular biologists in the manipulation of nucleic acids. One of the most important of these is Taq polymerase (Taq), which is a thermostable DNA polymerase named after the thermophilic bacterium Thermus aquaticus from which it was originally isolated. This enzyme is especially important, as it is central to the technique known as PCR, which allows sophisticated, targeted in vitro amplification and manipulation of sections of DNA or RNA. DNA

96


polymerases have also been central to the development of rapid modern methods of gene sequencing, which is one of the enabling technologies for the discipline of bioinformatics (interpretation of sequence information). Amongst the other enzymes which are important in molecular biology are those used to cleave DNA at specific sites (restriction endonucleases) and to join fragments of DNA (DNA ligases). Various other DNA-modifying enzymes are also used in vitro to help generate rDNA molecules (where DNA sequences from two or more sources which would not normally occur together are incorporated into a single ‘recombinant’ molecule). 2.3.1.3

The Genetic Code: Nucleotides, Codons and Amino Acids

In DNA molecules, the genetic code is represented by sequences of the four nucleotide bases adenine (A), cytosine (C), guanine (G) and thymine (T). On transcription, each template DNA base is represented in the equivalent mRNA by its complementary base; thus: DNA

!

RNA

Adenine Thymine Guanine Cytosine

! ! ! !

Uracil Adenine Cytosine Guanine

On the basis of this equivalence, if the DNA sequence is known, then the corresponding RNA sequence can be inferred, and vice versa. One of the basic units of genetic information in the genetic code is the codon, which is a specific tri-nucleotide sequence (triplet). There are four nucleotide bases (‘letters’) which can be arranged in three-letter combinations, making 64 possible codons (43 combinations) (see Figure 2.5). If the individual nucleotides can be thought of as the letters of the alphabet, then the codons resemble ‘words’, most of which ‘represent’ a corresponding amino acid. The exceptions are some codons called stop codons (UAG, UAA and UGA in RNA) which identify were a polypeptide ends, and the single start codon (AUG in RNA) which marks the start of a polypeptide-coding region in a sequence, and also corresponds to the amino acid methionine. Thus, identifying the codon sequence in coding DNA or mature mRNA can predict the polypeptide’s amino acid sequence. Only one grouping of the nucleotides into codons in a gene results in a correct amino acid sequence in the corresponding protein (this is referred to as the open reading frame). The loss or addition of a single nucleotide in a sequence causes a frameshift mutation, which results in a different sequence of codons (and, hence, a changed amino acid sequence). Frameshift mutations result in expression of altered polypeptide sequences, which are often truncated due to the premature occurrence of stop codons. To make things a little more complicated (or interesting), it is worth remembering that some amino acids are specified by more than one codon (termed redundancy; there are 20 basic amino acids and 64 available codons). Quite often, where an amino acid has associated codon redundancy, an organism will more frequently use one or more codons over the others to specify that particular amino acid (codon bias).

U

e r in se

U G

C

A

U

C

G

A A

A

C A G UC AG

U

C UC

A

isoleucine

hion

pr oli ne

ine

met

e

e

arginin

ine tam

ine

e

G U

U

G

G

C

leucin

tid

nin

STOP tryptophan

his glu

eo

A G U C AG UC

U

UC

G

ine

cyste

A

as

thr

OP ST

C

C

U

e

in

g ra pa

ine

G

ine

lys

U

C

e serin

G

U

arginine

97

os

tyr

G

G

valine

A

A

A

A GUCAG UC UC A G A

G

C A G UC A G U C GU

C

e

C

A

A

nin

phenyl alanine

glycine

ic tam glu d i c ac r ti pa as id ac

ala

leuc ine


Figure 2.5 The genetic code. The code has been illustrated by various formats: in tables, charts or, as shown here, as a wheel. This chart shows codons in RNA format (i.e. as they would be represented in mRNA). The nucleotides are represented in sequence by the three shaded concentric rings. The first nucleotide in the triplet is represented by the centre four sectors, the second by the next 16 sectors in the middle shaded ring, and the final position is designated by the outermost shaded ring. Some amino acids have unique codons (methionine AUG, tryptophan UGG), while others, such as arginine and threonine, are encoded by multiple codons (‘degeneracy’). Most organisms do not apply the code randomly when translating nucleic acids to proteins, but instead ‘prefer’ to use a subset of codons for some amino acids; this is called ‘codon bias’ and can sometimes cause difficulties in producing active enzymes from cloned DNA. The chosen host cell may have a codon bias which is not reflected in the cloned sequence; this can slow down or ‘stall’ the translation process and yield truncated or misfolded polypeptides (inactive enzyme). When this happens, various remedies can be adopted, including changing the host or changing the sequence artificially to suit the host’s codon preference (codon matching). The code illustrated here is often called the ‘universal code’; however, there are differences sometimes seen in various classes of organism, or by certain individual species, or even in different subcellular compartments of eukaryotic cells. It is recognized, for example, that human mitochondrial DNA uses a slightly different code from that found in human genomic DNA. Bioinformatics programs usually allow the researcher to specify which recognized version of the code is to be used when analysing DNA or RNA sequences to determine the likely associated protein sequence from coding regions

The assignment of amino acids to codons, sometimes called the universal genetic code, has been developed by comparison of many DNA and protein sequences. However, some organisms may routinely assign ‘unexpected’ amino acids to codons (nonstandard genetic code). Both codon bias and nonstandard genetic code can present problems when trying to get expression of genes in foreign hosts. As a general rule, for a cloned gene, if two nonstandard codons are contiguous then protein expression will be suppressed to very low levels.

98


Some DNA is referred to as noncoding, as it does not appear to specify a polypeptide. Large regions of eukaryotic genome comprise noncoding intergenic sequences and noncoding sequences which are present within gene sequences (such as introns). The protein-coding regions of the mouse and human genomes is about 3 % of the total genome. Prokaryotes typically have more ‘compact’ genomes, where protein-coding regions account for about 90 % of the genome. Other types of genetic information are specified by the noncoding regions – these are often related to regulatory functions, such as determining when the protein will be produced and how much protein will be produced in a given set of conditions. The term motif is often used for nucleic acid sequence specific information (both coding and noncoding regions). However, the function of much of the noncoding DNA in eukaryotes is not known, sometimes leading to it being referred to rather disparagingly as junk DNA. 2.3.1.4

Molecular Cloning and rDNA

When genetic information (nucleic acid) is transferred between different cells, species or genera it is often carried by a specialized DNA molecule called a vector. Viruses are natural vectors, as are some kinds of small independently replicating circular extrachromosomal DNA molecules (plasmids). A few of the basic features of plasmids used in molecular biology are reviewed in Figure 2.6. Commercial vectors are often derived from viruses or plasmids and now usually contain highly modified control systems for maintaining, amplifying or expressing gene sequences (‘cloned DNA’) in foreign host cells. Some vectors (shuttle plasmids) have been developed to be transferred between different kinds of host. This can be very useful when genetic manipulation is easy in one host (such as the laboratory favourite E. coli), but the functional protein (enzyme) can only be expressed in another type of system (such as in a yeast). Usually, the objective is to obtain a host cell which maintains the stability of the foreign gene. This is achieved either by using a vector which the host will continue to replicate separately from the host’s genomic (chromosomal) DNA or integrating the foreign DNA into the host’s genome at a target site by a process known as homologous recombination. When a stable genetic change has occurred in a cell due to the uptake of nucleic acid, this is often referred to as transformation. If a virus were involved in the process of transferring the information, this is sometimes termed transfection. rDNA is any form of DNA which has been produced by the combining of DNA sequences which would not be found together in nature. The resulting hybrid or chimeric DNA molecule is often simply called a construct. Organisms which contain rDNA molecules are referred to as genetically manipulated organisms. Usually, microbes are used for the production of enzymes using rDNA technology, and these then are sometimes called genetically modified microorganisms. Cloning simply refers to making many copies of something; informally, it can refer to rDNA technology which transfers copies of foreign genes to another DNA molecule and/or biological host cell (gene cloning). A clone usually refers to a subset of viable cells derived from a selection procedure to identify those individual cells which contain the desired rDNA construct. 2.3.1.5

Basic Gene Cloning from DNA Templates

Perhaps one of the simplest methods for obtaining an enzyme by rDNA technology is by using the well-established techniques of basic gene cloning, sometimes called shotgun cloning. The method is relatively simple but can be time consuming. Although many of the

2.3 Molecular Biology for Improved Biocatalysts Promoter

99

Multiple cloning site

ORI

PLASMID 1234bp

AntibioticR

Figure 2.6 Basic features of bacterial plasmids used in rDNA technology. An origin of replication (ORI) allows the plasmid DNA to be replicated by the host cell to ensure the plasmid’s propagation and survival. Plasmids can vary in copy number (number of plasmids per cell). The copy number can in turn affect the amount of product expressed by a gene dosage effect. The copy number of a vector can vary, and often the presence of a very large cloned insert can severely reduce the plasmid copy number. Selectable markers (such as drugresistance genes) are usually incorporated so that transformants can be selected on solid media and plasmids retained in culture. Controlled expression (production of protein from cloned gene) can be achieved using a vector promoter. The promoters are usually inducible, so that expression of the foreign gene is tightly controlled. This is very important in the early stages of a gene cloning experiment, as uncontrolled or excessive expression of a foreign gene product can sometimes be toxic for the host cell, resulting in loss of the clone. In order to be expressed under the control of the vector promoter, the cloned sequence has to be inserted in the correct orientation (as indicated by the arrowhead) usually with the start codon a short defined distance away from the promoter sequence. Multiple cloning sites allow accurate insertion of foreign DNA. The sites often contain several restriction sites (short target sequences recognized by restriction enzymes). Directional cloning is possible by using pairs of restriction sites to insert the foreign DNA in the required orientation (usually so it can be under the control of the vector promoter). Most plasmids can accept inserts in the range 1–10 kbp, although often clones with smaller inserts prove to be more stable than those containing large inserts. Where large fragments of DNA are to be cloned, then alternative vectors could include cosmids, yeast artificial chromosomes and bacterial artificial chromosomes. Other features often found in plasmids (or other vectors) include sites which allow direct cloning of PCR products or additional genes or sequences which can be fused to the foreign gene (to generate tagged proteins with additional amino acids, or even fuse the target gene with another enzyme). Some plasmids also have additional features, such as mechanisms for colorimetric detection of clones bearing inserts. ‘Suicide vectors’, which cannot be maintained autonomously in the cell, are also used to transform bacteria, yeasts and filamentous fungi. Plasmid components (including inserted genes) can only survive in transformants if they have successfully integrated into the host cell’s genome. ‘Shuttle vectors’ contain multiple origins of replication and selectable markers which allow them to be maintained in different hosts. This could be two bacterial species or bacteria plus yeast (or fungus). Copy number, selectable marker and promoter type are all important features to consider when choosing a plasmid for production of an enzyme

100


procedures have been superseded by other more modern techniques, it still serves to illustrate the basic cloning methods and is also still worth considering as a starting point in gene discovery. This is especially relevant where there is information on an enzyme’s activity but little or no sequence information available for the gene of interest. The DNA template in this case is total DNA from the original organism which produced the enzyme (for simplicity, preferably this will be a bacterium) (Figure 2.7). Usually, the bacterial cell walls are digested to release multiple copies of very high molecular weight DNA (genomic DNA). This genomic DNA is then broken up by physical shear force (passing through a syringe needle, for example), or digested by special enzymes which cut DNA at specific sites (restriction enzymes). The fragments are annealed (ligated) to a linearized vector DNA molecule using an enzyme from E. coli (DNA ligase). The ligation process can be more efficient if restriction enzymes have been used to generate the fragments and the cut vector, as this generates complementary short DNA single-strand ‘overhangs’ at the cut ends. These complementary ‘sticky ends’ can then anneal, stabilizing the hybrid construct prior to ligation. The population of hybrid molecules obtained is called a library. When the generation of inserted DNA fragments has been by random cutting or shearing it is sometimes called a shotgun library and the whole process is called shotgun cloning.

BASIC GENE CLONING Shotgun cloning experiment

Cell Physical shear

Genomic DNA

Enzymatic digest

Recombinant DNA Molecule

Vector

Enzymatic digest

Enzymatic ligation

Figure 2.7 Basic gene cloning I: shotgun cloning. Fragments of genomic DNA generated by physical shear or enzymatic digest (using restriction enzymes) are mixed with predigested plasmid vector. The ends of the linearized digested vector can be modified to reduce religation of the vector without insert. The religated vector (containing insert(s)) forms a library of rDNA molecules. Similar methods can be used to clone DNA fragments obtained by other methods – those recovered directly from the environment, sequences modified by mutagenesis, or cDNA generated from mRNA by reverse transcription


101

Genes from eukaryotes (yeast, fungi, plants and animals) and some Archaea pose some special cloning problems, as previously noted. The structure of genes in these organisms is fundamentally different from those in most bacteria. The coding sequences of eukaryote genes (‘exons’) are frequently interrupted by noncoding regions (called ‘introns’). Instead of using the technique of direct shotgun cloning of genomic DNA, the mature mRNA from these eukaryotic cells, which contains no introns, is converted to equivalent DNA molecules called complementary DNA (cDNA) in vitro using reverse transcriptase. Libraries of these cDNAs are then used in cloning experiments. The method has the advantage that only protein-coding regions are cloned (avoiding the cloning of introns and other noncoding DNA). On the downside, there is a bias towards cloning abundant transcripts (genes which are actively undergoing transcription when the mRNA was harvested). Poorly expressed genes might be difficult to detect, as there may be few or no copies of the particular mRNA present under some circumstances. Sometimes, a large proportion of transcripts are truncated and the upstream regulatory regions of the gene are not recovered in the cDNA cloning process. The DNA or cDNA library is then introduced into a preparation of bacterial host cells. Usually, the first host selected is a laboratory strain of E. coli which has been grown and pretreated with inorganic salts to make uptake of DNA easier. The ability to take up foreign DNA is called competence; cells which have been specially prepared for the purpose are called competent cells. Other methods to transfer DNA into cells include electroporation (application of an external electric field to permeabilize the cell wall), transfection (where a recombinant bacterial virus is used to transfer the DNA to the target cell) or ballistic methods (by using DNA-coated particle projectiles). The last method has been used to introduce foreign DNA into plant cells and mammalian cells. After a brief incubation of the competent cells in contact with the DNA library to allow uptake of the DNA, the bacterial cells are spread on Petri dishes containing sterile agar which usually also contains an antibiotic for selection. The objective of selection is to permit only those cells which have taken up the vector to grow; these cells are also more likely to contain ‘foreign’ cloned genes. A common method to select colonies of bacteria (clones) which contain the plasmid is to include a drug (antibiotic)-resistance marker on the plasmid vector and then add the antibiotic to the cell growth such that all cells without a plasmid are killed (Figure 2.8). Variations on this technique can be used to distinguish between clones with or without inserts in the vector, or to select for clones with DNA inserts of a defined size range. The colonies which grow on the agar are selected for individual culture (‘purification’) and subjected to various tests to check whether the gene of interest is present. The most direct form of test is for enzyme activity (functional test), but this could prove tedious, as a successful shotgun cloning can produce hundreds or thousands of colonies, each to be tested for the presence of functional enzyme. Where it is possible to include an enzyme assay directly (perhaps a colorimetric or fluorescence-based assay) in agar culture plate or multiwell format, the screening process can be significantly easier. These kinds of assay can be based on actual substrates reacting and inducing a measurable change (such as a pH change) or on related ‘artificial’ substrates which can emit a signal colour change or fluorescence when acted on by the enzyme. Artificial colorimetric substrates are attractive because a huge number of colonies can be screened relatively quickly, but they have the serious drawback that you are not directly selecting for the

102

Biocatalyst Identification and Scale-up: Molecular Biology for Chemists BASIC GENE CLONING Transformation and selection Transformation

+

Vector + insert

Competent host cells

(Library of recombinant DNA molecules)

Growth on selective medium

Bacterial colonies containing plasmids (library of transform)

Figure 2.8 Basic gene cloning II: transformation and selection. The library of rDNA molecules generated in vitro needs to be introduced into a suitable host by a process known as transformation. The uptake of DNA by the cells is enhanced by chemical treatments or by using electrical pulses. The cells now house the library. In order to distinguish between cells which do or do not contain the plasmid, most plasmids carry a selectable marker – a gene which modifies a characteristic of the host cells. Commonly used plasmids in basic cloning experiments often use antibiotic resistance as the marker. By incorporating the antibiotic into solid microbial growth medium (agar plates), only transformed cells containing the plasmid can grow into colonies. Some other useful features of plasmid biology ensure that each surviving cell in a single colony will have the same individual recombinant plasmid. Individual colonies are further analysed by functional tests of enzyme activity and/or by molecular screening for the required insert. The transformed cells are a convenient storage for the library and for any individual clone selected for further study. Cultures (individual or pooled) are often routinely stored at 70 C for several years without appreciable plasmid loss

activity of choice; and, as ‘you get what you screen for’, you may not be identifying the optimal clones at this stage. However, if the gene has been cloned, but the required activity is not produced, then the functional test will fail to pick up the target gene. In this case, if some gene sequence information is available, then it may be possible to test for the presence of DNA with the expected sequence by hybridization with radio-labelled ‘probe’ DNA or, more usually, by PCR. This sequence-based screening test could pick up ‘positives’ which have been missed in the initial screen because the gene has been successfully cloned but the enzyme has not been produced in an active form (perhaps because expression has not occurred or because E. coli is a poor host to support production of active enzyme), or where there is no convenient function-based assay available. Other methods include detection of production of mRNA or protein in response to the presence of the substrate (substrate-induced gene-expression screening). The success of this method is dependent on the retention of native upstream regulatory regions of genes in the clones which can switch on production of mRNA or enzyme in response to the stimulus of the presence of substrate. The method is used in screening libraries derived by ‘random’


103

cutting/shearing of genomic DNA into relatively large fragments, or where the cloned DNA fragments have been directly isolated from the environment. Both types of test could prove laborious, as many thousands of clones might be produced in a single experiment. However, there are several strategies available which could make the process simple, from using an initial selective growth test (functional test – determine conditions where survival of positive clone depends on production of active enzyme) to using pooled sub-libraries to reduce the number of sequence-based tests needed to identify a positive clone. Where a colorimetric or fluorescent-based assay is available it is also possible to use high-throughput screen methods based on cell sorting or automated colony picking. These facilities are expensive, but are available on a commercial basis for clone selection and are important when any high-throughput experiments are being considered. Some of the materials and techniques used in molecular biology may attract royalties if used for commercial purposes. Vectors, host strains and off-the-shelf DNA manipulation methods are usually readily available for modest licence fees for research purposes, but additional licences would need to be sought (and fees paid) if these systems were used in a commercial process. Where commercial exploitation is planned, the researchers should be prepared to switch to royalty-free genetic systems and avoid the use of costly and potentially toxic materials, such as artificial inducers or substrates, as gene expression regulators. 2.3.1.6

PCR

PCR (polymerase chain reaction) is a technique widely used in molecular biology. Its versatility as a technique for the manipulation of DNA and RNA sequences can probably not be exaggerated. In its simplest form, a chain reaction can be developed in which a DNA sequence template is exponentially amplified. PCR derives its name from one of its key components, a DNA polymerase used to amplify a target section of DNA by in vitro enzymatic replication. dsDNA recovered from genomic DNA or from vectors (plasmids) normally has two complementary strands that can be separated into single-stranded DNA (ssDNA) by heating (‘melting’) beyond the melting temperature Tm of the double-stranded form. When the DNA is cooled, eventually the complementary bases find each other and the double-stranded form is recovered (annealing). In PCR, short double-stranded sections of DNA are generated using synthetic oligomers (primers) complementary to the sequences flanking the target DNA sequence. The primers are typically 15–30 bases long and are designed to anneal to the ssDNA. The synthetic primers will also eventually form the termini of the amplified DNA segment. The DNA polymerase recognizes the partial dsDNA sequences formed on annealing of the primers to the single-stranded template and the enzyme initiates DNA replication, forming double-stranded strands using the exposed single strand as the template. As the PCR cycles progress, the short sections of DNA generated in previous cycles of replication are themselves used as templates in successive cycles. ‘Normal’ bacterial DNA polymerase would not be able to retain activity over the multiple temperature shifts of the reaction cycles, so the sustained reaction is made possible by the use of a thermostable DNA polymerase with a temperature optimum of about 70 C (such as Taq polymerase from the thermophilic bacterium T. aquaticus). Excess primer and deoxyribonucleotide triphosphates (dNTPs)

104


100

are also required in the reaction mix. Repeated cycles of denaturation, annealing and extension are required to generate a population of DNA fragments, all of which are copies of the target sequence flanked by the primer sequences. Early PCR experiments were performed by manual or robotic transfer of reaction vials between water baths or heating blocks. These have been superseded by several generations of programmable thermal cyclers. Successful PCR amplifies a single or a few copies of a target sequence of DNA by many orders of magnitude. The process is described schematically in Figure 2.9. The relatively simple basic format of PCR can be extensively modified to perform a wide array of genetic manipulations. The reaction products can be directly modified by altering the primer design (to introduce mutations or add ‘adaptamers’ to the ends of sequences, for example). Control of the reaction conditions and choice of polymerase enzymes can also alter the fidelity of the replication. The method can also be adapted to anneal sections of DNA which have overlapping sequences. The ability to amplify specific sequences based on a relatively small amount of sequence information (to enable primer design) means that DNA fragments can be easily amplified from many environments and from samples which may only have few copies of the target sequences – from genomic DNA, from purified plasmids, directly from cells and from environmental samples (e.g. clinical specimens, water, soil). As only a few copies of the target sequence are required, the method is very sensitive, making it useful in amplification of sequences from mixed populations (as in metagenomics) or for molecular fingerprinting (speciation, genotyping and forensic applications). Careful experimental design can also modify the sensitivity and

Denature DNA

Temperature 50

Extend primers

Anneal Primers

0

Repeat n times .

Figure 2.9 Schematic representation of a typical simple PCR reaction. The starting reaction mix contains dsDNA template, a pair of short ssDNA oligonucleotide primers (complimentary to ends of target DNA sequence), a pool of the four dNTPs and a heat-resistant DNA polymerase, e.g. Taq polymerase. dsDNA containing the target sequence is denatured to the singlestranded form by heating to 90–100 C. The reaction is cooled to a few degrees below the calculated annealing temperatures of the synthetic primers to the target DNA sequence. At the extension temperature (often 72 C), Taq polymerase initiates the extension of the partial dsDNA segment using the single strand as a template. Successive denaturation, primer annealing and extensions produces copies of the target. The process is repeated n times (typically n ¼ 20–30 for a simple amplification experiment). Amplification factor: 2n


105

specificity of PCR. The versatility of the basic PCR technique in manipulation and analysis of DNA make it central to many molecular biology procedures. Many innovations in PCR technique have increased it versatility in identifying and recovering enzyme genes. For example, it is possible to devise PCR-based methods to recover genes based on limited information relating to short internal gene DNA sequences, or by using amino acid sequences from purified proteins to predict and detect the actual DNA sequence (this technique is sometimes called ‘reverse genetics’). Bioinformatics tools can be used to generate ‘consensus sequences’ for the active site of specific families of enzymes and the target DNA can then be probed for the presence of these sequences. One of the most problematic steps encountered when generating libraries of DNA sequences is often the enzymatic ligation of the DNA into vectors. Ligation can be the most inefficient step in the entire process, and the enzymes (ligase) and cofactors required are also relatively expensive. As a result, some of the diversity and complexity of the library can easily be lost at this stage. Short (usually two to four nucleotides) complementary single-strand overhangs are generated by restriction enzymes. When these are annealed together the hybrid molecules require in vitro stabilization by DNA ligase prior to transformation of the microbial host. Ligation-independent cloning methods based on modified PCR techniques generate complementary long sticky ends of 12–15 nucleotides on both the plasmid and the insert. The annealed insert/vector molecules generated are sufficiently stable to be transformed directly into E. coli, where the DNA backbone can be efficiently repaired by ligases in the host cell.25 2.3.2 2.3.2.1

Mutagenesis Directed Evolution of Enzymes

The huge diversity of enzymes observed in nature is attributed to evolutionary processes where diverse populations of gene variants (with sequence variations generated by mutation and recombination) are subjected to selection of the ‘fittest’ enzyme functions. Those genes which confer advantageous traits to their hosts are more likely to be maintained and disseminated in populations than those that do not. This process can be mimicked experimentally to modify enzymes by generating or enhancing enzyme gene diversity via mutation and recombination and then devising specific selection methods to identify ‘improved’ versions of the enzyme, which are then amplified for further analysis and manipulation. The diversification, selection and amplification can be thought of as the basic processes of directed evolution. Directed evolution provides a powerful tool for the development of biocatalysts with novel properties, without requiring knowledge of enzyme structures or catalytic mechanism.26 The initial step involves the identification and isolation of a ‘wild-type’ naturally occurring gene responsible for encoding the desired enzyme. This requires the cloning of the relevant gene into an efficient expression system before this target gene is subjected to random (or rational) mutagenesis using the methods such as those described in these sections. Improved variants are identified through screening of the function of the expressed enzymes (preferably by a carefully designed high-throughput screening method). The inferior enzymes and their genes are discarded and the improved genes are used as parents for the next round of evolution by repeating the diversification and selection process as

106

Biocatalyst Identification and Scale-up: Molecular Biology for Chemists Bacterial transformation

Random mutagenesis

Parental Genes

Error prone pcr Mutator strains recombination

Mutant library

Repeat to generate Improved enzyme variants

Screening and selection for improved enzyme function 1St Generation mutated gene

Figure 2.10 Generation of improved enzymes by directed evolution. The process starts with one or more parental genes which are subjected to mutagenesis – either by generation of random point mutations or small insertions/deletions, and/or by recombination of gene fragments to generate libraries of mutants. In the next stage this library is screened for the desired enzyme function. Ideally, this is combined with a selection or discrimination process as part of a high-throughput process. In the third stage, the selected mutants are isolated and amplified (by propagation of the expression clone or by PCR). These selected first-generation mutants then undergo successive rounds of directed evolution, each time selecting for the favourable features in the expressed enzymes. There are many variations on the technique: sometimes, selected gene domains (cassettes) are targeted for mutagenesis; iterative processes could be used in the generation of the mutant library, and specific sequence information may be used in the targeting of the mutagenesis or in the screening protocols

often as necessary. The basis steps of the process are summarized in Figure 2.10. Applying different techniques has resulted in the development of enzymes with improved properties and production of ‘new’ enzymes with diverse properties, such as improved enantioselectivity, activity, thermostability, protein solubility and expression. 2.3.2.2

Error-prone PCR

One of the commonest methods to achieve gene diversity by random mutagenesis is errorprone PCR (epPCR). This technique exploits the fact that the thermostable polymerase used for PCR has relatively low replication fidelity. It lacks a 30 to 50 exonuclease proofreading activity mechanism to replace any accidental mismatch in the newly synthesized DNA strand and has an error rate measured at about 1 in 10 000 nucleotides. To enhance this mutagenic effect, protocols have been developed with the aim of deliberately increasing the error rate of Taq polymerase, which can be varied by increasing the concentration of MgCl2, by addition of MnCl2 or by using unbalanced dNTP concentrations in the reaction mix to achieve higher rates of mutations. Point mutations are the most common types of mutation in epPCR, but deletions and frameshift mutations are also possible, although rarer. Other ways of increasing the mutation rate can include the use of natural or proprietary polymerases with enhanced mutation frequency27 and by incorporating synthetic mutagenic dNTPs, such as 8-oxo-dGTP, which are then eliminated in a subsequent


107

PCR reaction using natural dNTPs.28 However, there are some drawbacks to epPCR. Generally, the technique produces libraries of DNA fragments which need to be ligated into expression plasmids (this can be a limiting step), and some of the methods do not produce random mutations. 2.3.2.3

Cassette Mutagenesis and Mutator Strains

Another approach to mutagenesis is to restrict the mutagenesis of the gene to defined regions or ‘cassettes’. These may have been targeted as key domains by other bioinformatics or mutagenesis studies. Typically, a target region is excised by cutting the DNA at two constructed or naturally occurring restriction enzyme sites that flank this region and the excised portion is replaced with oligonucleotide(s) containing the desired mutation. The mutagenesis target could be as small as a single codon (thus, changing a single amino acid), and the type and degree of mutagenesis can be varied depending on the technique used (epPCR, synthetic oligonucleotides or use of in vivo mutagenesis techniques, such as mutator strains). Complete saturation of one or more positions in a protein with all possible amino acid substitutions can be achieved with this method. The technique has the disadvantage of being relatively expensive if large quantities of synthetic nucleotides are required; and, depending on the methodology employed, ligation steps may be required to recover the reconstructed gene. It can also be used in directedevolution experiments to generate libraries of genes which contain both conserved and highly mutated domains. Other useful random mutagenesis methods are based on introducing the target DNA into a host cell which has error-prone DNA replication processes. The most popular of these is probably the mutator strain method. Commercial strains, such as the E. coli XL1-Red (Stratagene, La Jolla, CA), lack three of the primary DNA repair pathways, MutS, MutD and MutT, resulting in a random mutation rate 5000-fold higher than in wild type. The protocol for using the mutator strain is composed of two steps: transformation of the mutator strain and recovery of the mutant from the transformant. In some ways the technique is much simpler than epPCR, and ligation steps are eliminated as the mutated gene is recovered in a plasmid vector. However, the actual mutation frequency can be fairly low under the standard conditions (0.5 mutations per kilobase), and extended cultivation periods are often required to introduce multiple mutations. 2.3.2.4

DNA Shuffling

In studying the mechanisms of gene evolution it is important to recognize the importance of recombination of blocks of sequence, rather than point mutagenesis alone, in generating sequence (and function) diversity. The DNA shuffling approach involves mixing of a family of homologous sequences obtained from nature (typically the same gene from related species or related genes from a single species) or from libraries of artificially mutated genes which creates a large diversity of novel structure–function proteins. The method shares some of the features of natural genetic recombination, in that new genes are generated by combining sections from the ‘parental’ genes. A basic feature of gene shuffling is that genes or fragments of genes are cut into fragments and reassembled as chimeric molecules. The library of chimeric sequences is inserted into expression plasmids and screened for desirable traits. The development of DNA shuffling by Stemmer in 1994

108


Starting genes

DNase 1 Gene fragments

Extend

Denature and anneal

Extend ‘Shuffled’ gene library

Figure 2.11 Gene shuffling to generate chimeric gene libraries. A ‘classical’ DNA-shuffling strategy begins by fragmenting a pool of double-stranded parent genes randomly using partial enzymatic digest with DNase I. A further refinement can be included by selection of small fragments by size fractionation to maximize the probability of multiple recombination events occurring. The fragments are recombined in vitro by allowing annealing of homologous sequences. The short sections of dsDNA formed then can act in a similar way to conventional PCR primers, allowing the fragments to ‘cross-prime’ each other in a round of primerindependent PCR amplification. Successive rounds of product annealing and amplification generate a library of ‘shuffled’ genes. In order to facilitate expression cloning of the gene library, the full-length, diversified products are usually then modified by an additional round of PCR amplification with terminal primers to allow insertion of the sequences into expression vectors

overcame some of the various drawbacks of random mutagenesis alone, in that a much greater diversity of useful mutants could be generated by making chimeric genes.29 A simplified representation of DNA shuffling is given in Figure 2.11. 2.3.2.5

Combinatorial Methods

There has been a rapid expansion in the past decade in methods for creating libraries using directed evolution by gene mixing techniques. Many combinations of mutagenesis and shuffling have been developed. More sophisticated approaches have also included application of statistical analysis of protein sequence–activity relationships to identify beneficial mutations in early round variants (including variants with reduced activity) and then combining these mutations by the incorporation of synthetic oligonucleotides with DNA


109

shuffling. A multitude of techniques have been described based on combinations of procedures to generate mutations, recombine gene fragments and isolate improved mutants. Techniques include: staggered extension protocol, (StEP), iterative truncation for the creation of hybrid enzymes (ITCHY), degenerate oligonucleotide gene shuffling (DOGS), sequence homology-independent protein recombination (SHIPREC), random chimeragenesis on transient template (RACHITT), synthetic shuffling, sequenceindependent site-directed chimeragenesis (SISDC), combination libraries enhanced by recombination in yeast (CLERY) and THIO-ITCHY to name a few. Several methods have been patented and the processes commercialized. We recommend the interested reader to consult more specialist literature to get a deeper understanding of these techniques. See Ref. 30, for example, for further reading. 2.3.2.6

Emerging Methods in Directed Evolution: Neutral Drift and Indels

In directed evolution the target mutation rates are orders of magnitude higher than those of nature (greater than one mutation per gene per generation from approximately 1 in every 106 in most natural organisms). Enzymes tolerate most single mutations with no loss of function, but their stability and the ability to tolerate more mutations is often severely compromised. Enzymes used as starting points for laboratory evolution were never evolved to withstand high mutational loads. Using more-stable enzymes (perhaps those from thermophiles) or laboratory-evolved stabilized enzymes is predicted to give better quality libraries. Another way is the neutral drift technique, where a starting library is first evolved with mutations selected to maintain the protein’s original function; this has been shown to be a way of generating small and highly effective libraries for directed evolution.31 Neutral-drift library sequence analysis has suggested that these mutations act by enriching ‘global suppressor’ mutations which increase the enzyme stability and suppress the effect of a broad range of otherwise destabilizing mutations. These mutations often involve sequence changes which result in drift to ‘back-to-consensus’/ ‘ancestral enzyme’ sequences. All these techniques create genetic diversity by recombination and point mutations and are well developed. However, insertions and deletions (indels) are also important types of mutation which are probably underrepresented in many conventional mutagenesis strategies. Methods for incorporation of indels in predefined positions in a combinatorial manner have been developed.32 Although there are some published studies on their use in the directed evolution of biocatalysts,33 the full potential of these newer methods of gene mutation for enzyme improvement are yet to be demonstrated. 2.3.2.7

Rational Enzyme (Re)design

The application of random mutagenesis or recombinatorial DNA shuffling methods to genes coupled with screening and selection has frequently been successfully applied to generate mutated enzymes with improved characteristics. These methods often do not require any specific knowledge of the enzymes’ tertiary structure. However, a completely ‘random’ mutagenesis approach can generate huge numbers of variants to be screened in directed-evolution experiments. Other approaches use a rational or semi-rational technique to ‘target’ mutations to subsections of sequence or individual codons which are associated with critical amino acids, such as those in or near the enzyme active site, for

110


example. In several instances, critical amino acid changes which can alter enzyme performance have been first identified by random mutagenesis strategies. Often these amino acids are at locations which are distant from the active site and their importance would not have been inferred by knowledge of the protein’s structure. The role of these distal residues in enzyme activity could then be further investigated by saturation mutagenesis at these sites.34 However, where changes in the enzyme’s substrate specificity or kinetics are sought, investigators often target the active site residues. The modification of these limited numbers of residues generates much smaller libraries of mutants. Individual amino acid changes can be investigated, or, as has been recently described, combinations of relevant residues can be simultaneously mutated.35 In a technique termed ‘CASTing’ (combinatorial active site saturation test), small subsets of active site residues (typically three residues) are subjected to saturation mutagenesis (where every possible protein amino acid is substituted for the native one). Beneficial mutations are re-entered into successive rounds of iterative saturation mutagenesis based on the same principles, with selection for improved performance. By this means, several small catalytically diverse libraries can readily be generated and subjected to successive rounds of directed evolution. Recent advances in enzyme engineering have used a combination of the more ‘random’ methods of directed evolution with elements of rational enzyme modification to try to overcome the limitations of both directed evolution (very large libraries, hard to screen) and rational design (based on knowledge of enzyme tertiary structure, which is frequently limited).36 Semi-rational approaches that target multiple, specific residues to mutate on the basis of prior structural or functional knowledge have the potential to create ‘smart’ libraries that are more likely to yield positive results. Emerging techniques combine bioinformatics to model protein sequence–function relationships and provide a rational basis for identifying ‘beneficial diversity’ to be investigated in further rounds of enzyme evolution.37 2.3.3

Gene Synthesis

The entire process of identifying a gene based on sequence information, using PCR or shotgun cloning to recover the entire sequence from DNA to generate a construct suitable for expression and troubleshooting expression, can be a time-consuming and expensive process. The increased availability of commercial oligonucleotide synthesis and sequencing facilities has accelerated the pace of molecular biology research significantly in the past decade, and this has recently been supplemented by the availability of relatively lowcost gene synthesis services. No intact template DNA is required in this case, but it is necessary to have actual sequence information (perhaps from a search of existing databases or as a result of sequencing experiments) or a novel ‘hypothetical’ sequence based on bioinformatics. 2.3.4

Overcoming Problems of Codon Bias

A good example where access to synthetic gene technology superseded laborious conventional genetic manipulation is demonstrated in the case of Candida rugosa lipase 1 (lip1).38 C. rugosa (also known as Candida cylindracea) is a yeast which produces a mixture of lipases (known as isoforms). In order to get access to a single pure lipase (lip1) in significant quantities it was necessary to isolate the gene for this enzyme, clone and

2.4 Microbiology and Fermentation

111

express it in a suitable host. Unfortunately, C. rugosa has unconventional codon usage and, more unfortunately, this yeast frequently uses the codon CUG to represent serine,39 even in the Ser209 at the catalytic site. In most organisms this codon represents leucine; so, when the unmodified lip1 gene was cloned and transferred into S. cerevisiae, an inactive enzyme was produced – presumably because 17 out of the total 47 serines (including that at the active site) had been replaced by leucine residues. The options for changing the sequence of the gene to introduce serine codons which would be correctly translated by S. cerevisiae or most other conventional hosts were (i) the laborious site-directed mutagenesis of the individual serine codons in the cloned gene until active enzyme could be produced or (ii) the generation of a new completely codon-optimized sequence by splicing synthetic oligomers together to reconstruct the 1.7 kbp gene. Option (ii) proved to be the more successful, not least because the ‘synthetic gene’ approach allowed several other modifications to be included which assisted in the cloning and expression of the enzyme (codon optimization to match the host preference, removal of unwanted restriction sites). A decade on, option (ii) would prove even more rapid and less expensive, as commercial gene synthesis has become widely available. Provided with DNA or protein sequence, commercial gene synthesis companies can rapidly generate the complete sequence as dsDNA. The sequence can be optimized to overcome codon bias and can be supplied inserted in an appropriate vector for the selected expression host – bacterial, fungal, insect or mammalian. Site-specific mutagenesis is usually also an option during the gene synthesis (by incorporating a range of nucleotides at any specific site or sites), so a ‘library’ of specific mutants can also be supplied at the outset. At the time of writing, a ‘typical’ enzyme gene of 2–3 kbp could be produced in an expression-ready vector in a matter of days or weeks for under US$5000.

2.4 2.4.1

Microbiology and Fermentation E. coli as an Expression Host: the Pros and Cons

E. coli is often the first choice as host for many research purposes due to the large amount of commercial vector, transformation and expression systems available and because gene expression is relatively well understood in this organism. It is also easy to cultivate, with visible colonies growing from single cells in less than 24 h. However, E. coli frequently presents several problems when it comes to producing large quantities of functional purified enzyme. Foreign proteins are usually not exported by E. coli, so in order to obtain purified enzyme it may be necessary to lyse the cells by using detergents and/or physical shock (sonication, shear stress). Inside the cell, overexpressed proteins may have accumulated in insoluble aggregates (inclusion bodies). Often, the inclusion bodies contain misfolded (inactive) protein. Various strategies can be employed to overcome the limitations of E. coli as an expression host. These include increasing the copy number of the vector (and, therefore, increasing the number of copies of the cloned gene per cell) and using various induction methods and cell culture strategies to increase expression of functional protein. Occasionally, the expressed foreign protein is itself toxic for the host, so expression must be very carefully controlled. Various strategies can be adopted to overcome these

112


problems, including reducing the rate of production of proteins (lowering temperature, using stricter induction controls). Proprietary strains of E. coli have also been developed which have been genetically modified to overcome codon bias, promote correct folding, post-translational modification and promote secretion of active enzyme.40 Inactive enzyme derived from purified inclusion bodies can sometimes be refolded in vitro to produce active enzyme. It may also be important to deal with issues of codon bias or nonstandard genetic code, if necessary by codon matching of the introduced sequence (substituting mutated or synthesized gene sequences with the appropriate ‘host’ codons for those found in the ‘native’ sequence). In addition, there are problems sometimes encountered when trying to maintain plasmids in scaled-up or extended cultures. These can be due to the need to avoid the use of allergenic antibiotics and/or inherent plasmid instability (often accompanied by multimerization of the plasmids). These issues can sometimes be overcome by use of specialized hosts, by careful selection of culture conditions to optimize plasmid retention or by adoption of alternative selection mechanisms; for example, those based on complementing auxotrophy. Auxotrophy is the inability of an organism to synthesize a particular organic compound (often an amino acid or vitamin) required for its growth. By replacing the missing gene on the transforming expression plasmid, the autotrophic cell acquires the ability to grow in the absence of the amino acid or vitamin.41 2.4.2

Alternative Host Expression Systems: Bacteria, Yeasts, Filamentous Fungi

Alternative hosts are sometimes chosen for their ability to produce secreted products (E. coli cells tend to produce intracellular proteins), to improve the probability of ‘correct’ translation and to incorporate post-translational processing of polypeptides (such as glycosylation, folding, cross-linking and export). However, initial genetic manipulations are often carried out in E. coli and the relevant construct then transferred into the ‘production’ host. For certain applications, such as when the product may be used in pharmaceutical, food, feed or personal care markets, it may be preferable to use hosts which are not associated with pathogenicity or toxicity (‘generally recognized as safe’– examples include S. cerevisiae and Bacillus subtilis). Where large quantities of recombinant enzyme are required, successive rounds of cloning experiments may be carried out to place the gene in the appropriate vector, design a controlled expression system and choose a host cell which can be grown on the appropriate industrial scale. In order to overcome some of the difficulties associated with heterologous gene expression in E. coli, various alternative microbial hosts have been developed (e.g. Bacillus, Pseudomonas (DowPharma PfenexTM Expression Technology)). Various yeasts (S. cerevisiae, Hansenula polymophia, Pichia pastoris, Yarrowia lipotytica) are frequently used as an expression system for the production of proteins. A number of properties that make Pichia suited for this task include its high growth rates and an ability to grow on a simple, inexpensive medium. Pichia can be readily grown in either shake flasks or fermenters, which makes it suitable for both small- and large-scale production. Filamentous fungi such as Aspergillus species are also excellent hosts for the production and export of enzymes. Although their genetic manipulation is initially a bit more time consuming compared with yeasts or bacteria, they readily form stable transformants which can be grown at large scale for industrial enzyme production.42

References

2.5

113

Summary, Overview and Future

While reactions catalysed by ‘wild-type’ enzymes will remain a basic tool kit for biocatalysis, the increased accessibility of gene manipulation (expression optimization, functional optimization by mutagenesis) will make lower cost ‘bespoke’ enzymes more readily available. Not only will a wider range of enzymes be available in the future, but the lead times in process development may also be reduced as the catalyst (the enzymes and/or the cell) can be more rapidly modified to suit the desired process conditions. Advances in bioinformatics and improvements in reaction modelling could mean that, from biocatalyst screening (discovery) right through to enzyme optimization, process scale-up could be contracted into a few weeks. In conjunction with improvements in enzyme function, there have been significant recent advances in improving host cells. Progress has already been made in the redesign of microbial metabolic networks to generate strains optimized for production of small molecules.43 Other research is focusing on the radical rational ‘stripping down’ of microbial genomes to generate ‘simplified’ cells with the minimum number of characterized functional genes – usually with the aim of enhancing protein (enzyme) production by removing bottlenecks. These ‘minimum genome factories’ have been proposed for various platform biotechnology strains, including those of E .coli and B. subtilis.44 Further advancements in systems biology (advanced studies of complex biological systems) could in the future help to establish ‘synthetic biology’ programmes – construction and design of artificial biological system parts or whole organisms. Delivery of these technological advances will depend on the multidisciplinary efforts of scientists and engineers, but could produce reliable and efficient commodity and biocatalyst manufacturing platforms of the future.

References 1. Peterson, D.H., Murray, H.C., Eppstein, S.H., Reineke, L.M., Weintraub, A., Meister, P.D. and Leigh, H.M., Microbiological transformations of steroids. I. Introduction of oxygen at carbon-11 of progesterone. J. Am. Chem. Soc., 1952, 74, 5933–5936. 2. Woodward, R.B., Sondheimer, F., Taub, D., Heusler, K. and McLamore, W.M., The total synthesis of steroids. J. Am. Chem. Soc., 1952, 74, 4223–4251. 3. Roberts, S.M. and Wiletts, A.J., Introduction to Biocatalysis Using Enzymes and Microorganisms, 2nd edition. Cambridge University Press, 1995. 4. Savile, C.K, Magloire, V.P. and Kazlauskas, R.J., Subtilisin-catalyzed resolution of N-acyl arylsulfinamides. J. Am. Chem. Soc., 2005, 127, 2104–2113. Dawson, M.J., Mahmoudian, M. and Wallis, C.J., Process for preparing enantiomerically enriched N-derivatised lactams. 1999, WO/1999/010519. 5. Schomburg I., Chang A., Hofmann O., Ebeling C., Ehrentreich F. and Schomburg, D., BRENDA: a resource for enzyme data and metabolic information. Trends Biochem. Sci., 2002, 27, 54–56. 6. Ellis L.B.M., Roe, D. and Wackett, L.P., The University of Minnesota Biocatalysis/ Biodegradation Database: the first decade. Nucleic Acids Res., 2006, 34, 517–521. 7. Altschul, S.F., Gish, W., Miller, W., Myers, E.W. and Lipman, D.J., Basic local alignment search tool. J. Mol. Biol., 1990, 215, 403–410. 8. Sigrist C.J.A., Cerutti L., Hulo N., Gattiker A., Falquet L., Pagni M., Bairoch A. and Bucher, P., PROSITE: a documented database using patterns and profiles as motif descriptors. Brief Bioinform., 2002, 3, 265–274.

114


9. Handelsman, J., Rondon, M.R., Brady, S.F., Clardy, J. and Goodman, R.M., Molecular biological access to the chemistry of unknown soil microbes: a new frontier for natural products. Chem. Biol., 1998, 5, 245–249. Handelsman, J., Metagenomics: application of genomics to uncultured microorganisms. Microbiol. Mol. Biol. Rev., 2004, 68, 669–685. 10. Ferrer M., Golyshina O., Beloqui A. and Golyshin P.N., Mining enzymes from extreme environments. Curr. Opin. Microbiol., 2007, 10, 207–214; Liu J.R., Yu B., Lin S.H., Cheng K.J. and Chen Y.C., Direct cloning of a xylanase gene from the mixed genomic DNA of rumen fungi and its expression in intestinal Lactobacillus reuteri. FEMS Microbiol. Lett., 2005, 251, 233–241. 11. Bull, A.T., Microbial Diversity and Bioprospecting. ASM Press, 2004. 12. Yun, J. and Ryu, S., Screening for novel enzymes from metagenome and SIGEX, as a way to improve it. Microb. Cell Fact., 2005, 4, 8.Uchiyama, T., Abe, T., Ikemura, T. and Watanabe, K., Substrate-induced gene-expression screening of environmental metagenome libraries for isolation of catabolic genes. Nat. Biotechnol., 2005, 23, 88–93. 13. Ji, S.C., Dockyu, K., Jung-Hoon,Y., Oh, T.-K. and Choong-Hwan,L., Sequence-based screening for putative polyketide synthase gene-harboring clones from a soil metagenome library. J. Microbiol. Biotechnol., 2006, 16, 153–157; Ginolhac, A., Jarrin, C., Gillet, B., Robe, P., Pujic, P., Tuphile, K., Bertrand, H., Vogel, T.M., Perrie`re, G., Simonet, P. and Nalin, R., Phylogenetic analysis of polyketide synthase I domains from soil metagenomic libraries allows selection of promising clones. Appl. Environ. Microbiol., 2004, 70, 5522–5527. 14. Gabor, E.M., de Vries, E.J. and Janssen, D.B., Construction, characterization, and use of smallinsert gene banks of DNA isolated from soil and enrichment cultures for the recovery of novel amidases. Environmental Microbiol., 2004, 6, 948–958. 15. Littlechild, J.A., Guy, J., Connelly, S., Mallett, L., Waddell, S., Rye, C.A., Line, K. and Isupov, M., Natural methods of protein stabilization: thermostable biocatalysts. Biochem. Soc. Trans., 2007, 35, 1558–1563. 16. Dalmau, E., Montesinos, J.L., Lotti, M. and Casas, C., Effect of different carbon sources on lipase production by Candida rugosa. Enzyme Microb. Technol., 2000, 26, 657–663. 17. Gonzalez-Lopez,C., Szabo, R., Blanchin-Rolanda,S. and Gaillardina, C., Genetic control of extracellular protease synthesis in the yeast Yarrowia lipolytica. Genetics, 2002, 160, 417–427. Voigt, B., ThiHoi, L., Ju¨rgen,B., Albrecht, D., Ehrenreich, A., Veith, B., Evers, S., Maurer, K.H., Hecker, M. and Schweder, T., The glucose and nitrogen starvation response of Bacillus licheniformis. Proteomics, 2007, 7, 413–423. 18. Better, M.D., Improved methods and bacterial cells for expression of recombinant protein products. PCT Int. Appl., 2001, WO 2001073082. 19. Chater, K.F., Bruton, C.J., O’Rouke, S.J. and Wietzorrek, A.W., Methods and materials relating to gene expression. 2002, EP1244799 (A1). 20. Hermanna, M., Kietzmanna, M.U., Ivancˇicá, M., Zenzmaiera, C., Luitenb, R.G.M., Skrancc, W., Wubbolts, M., Winklere, M., Birner-Gruenbergerf, R., Pichlera, H. and Schwab, H., Alternative pig liver esterase (APLE) – cloning, identification and functional expression in Pichia pastoris of a versatile new biocatalyst. J. Biotechnol., 2008, 133, 301–310. 21. Osprian, I., Fechter, M.H. and Griengl, H., Biocatalytic hydrolysis of cyanohydrins: an efficient approach to enantiopure -hydroxy carboxylic acids. J. Mol. Catal. B: Enzymatic, 2003, 24–25, 89–98. 22. Reisinger, C., Osprian, I., Glieder, A., Schoemaker, H.E, Griengl, H. and Schwab, H., Enzymatic hydrolysis of cyanohydrins with recombinant nitrile hydratase and amidase from Rhodococcus erythropolis. Biotechnol. Lett., 2005, 26, 1675–1680. 23. Burton, S.G., Cowan, D.A. and Woodley, J.M., The search for the ideal biocatalyst. Nat. Biotechnol., 2002, 20, 37–45. 24. Allison, L.A. Fundamental Molecular Biology, Wiley-Blackwell, 2006. Kreuzer, H. and Massey, A., Molecular Biology and Biotechnology: A Guide for Students, 3rd edition. John Wiley & Sons, Ltd, 2008. 25. Aslanidis, C. and de Jong, P.J., Ligation-independent cloning of PCR products (LIC-PCR). Nucleic Acids Res., 1990, 18, 6069–6074. 26. Turner, N., Directed evolution of enzymes for applied biocatalysis Trends Biotechnol., 2003, 21, 474–478; Johannes, T.W. and Zhao, H., Directed evolution of enzymes and biosynthetic pathways. Curr. Opin. Microbiol., 2006, 9, 261–267.

References

115

27. Examples of proprietary enzymes include Mutazyme II DNA Polymerase (www. stratagene.com). Other companies have developed ‘kits’ containing selections of reagents which can be combined to yield controlled error rates in PCR reactions. 28. Zaccolo, M., Williams, D.M., Brown D.M. and Gherardi, E., An approach to random mutagenesis of DNA using mixtures of triphosphate derivatives of nucleoside analogues. J. Mol. Biol., 1996, 255, 589–603. 29. Stemmer, W.P., DNA shuffling by random fragmentation and reassembly: in vitro recombination for molecular evolution. Proc. Natl. Acad. Sci. U. S. A., 1994, 91, 10747–10751. 30. Arnold, F.H. and Volkov, A.A., Directed evolution of biocatalysts. Curr. Opin. Chem. Biol., 1999, 3, 54–59. Thomas, J.M. and Raja, R., Designing catalysts for clean technology, green chemistry, and sustainable development. Annu. Rev. Mater. Res., 2005, 35, 315–350. Kaur, J. and Sharma, R., Directed evolution: an approach to engineer enzymes. Crit. Rev. Biotechnol., 2006, 26, 165–199. 31. Gupta, R.D. and Tawfik, D.S., Directed enzyme evolution via small and effective neutral drift libraries. Nat. Methods, 2008, 5, 939–942. 32. Fujii, R., Kitaoka, M. and Hayashi, K., RAISE: a simple and novel method of generating random insertion and deletion mutations. Nucleic Acids Res., 2006, 34, 30. 33. Bershtein, S. and Tawfik, D.S., Advances in laboratory evolution of enzymes. Curr. Opin. Chem. Biol., 2008, 12, 151–158. 34. Parikh, M.R. and Matyoumara, I., Site-saturation mutagenesis is more efficient than DNA shuffling for the directed evolution of -fucosidase. J. Mol. Biol., 2005, 352, 621–628. 35. Reetz, T., Wang, L.-W. and Bocola, M., Directed evolution of enantioselective enzymes: iterative cycles of CASTing for probing protein sequence space. Angew. Chem. Int. Ed., 2006, 45, 1236–1241. 36. Chica1, R.A., Doucet, N. and Pelletier, J.N., Semi-rational approaches to engineering enzyme activity: combining the benefits of directed evolution and rational design. Curr. Opin. Biotechnol., 2005, 16, 378–384. 37. Fox, R.J. and. Huisman, G.W., Enzyme optimization: moving from blind evolution to statistical exploration of sequence–function space. Trends Biotechnol., 2008, 26, 132–138. 38. Brocca, S., Schmidt-Dannert, C., Lotti, M., Alberghina, L. and Schmid, R.D., Design, total synthesis, and functional overexpression of the Candida rugosa lip1 gene coding for a major industrial lipase. Protein Sci., 1998, 7, 1415–1422. 39. Kawaguchi, Y., Honda, H., Taniguchi-Morimura,J. and Iwasaki, S., The codon CUG is read as serine in an asporogenic yeast Candida cylindracea. Nature, 1989, 341, 164–166. 40. Choi, J.H. and Lee, S.Y., Secretory and extracellular production of recombinant proteins using Escherichia coli. Appl. Microbiol. Biotechnol., 2004, 64, 625–635. 41. Vidal, L., Pinsach, J., Striedner, G., Caminal, G. and Ferrer, P. Development of an antibiotic-free plasmid selection system based on glycine auxotrophy for recombinant protein overproduction in Escherichia coli. Biotechnol., 2008, 134, 127–36. 42. Punt, P., van Biezen, N., Conesa, A., Albers, A., Mangnus, J. and van den Hondel, C., Filamentous fungi as cell factories for heterologous protein production. Trends Biotechnol., 2002, 20, 200–206. 43. Pharkya, P., Burgard, A.P. and Maranas, C.D., OptStrain: a computational framework for redesign of microbial production systems. Genome Res., 2004, 14, 2367–2376. 44. Ara, K., Ozaki, K., Nakamura, K., Yamane, K., Sekiguchi, J. and Ogasawara, N., Bacillus minimum genome factory: effective utilization of microbial genome information. Biotechnol. Appl. Biochem., 2007, 46, 169–178; Mizoguchi, H., Mori, H. and Fujio, T., Escherichia coli minimum genome factory. Biotechnol. Appl. Biochem., 2007, 46, 157–167; Mizoguchi, H., Mori, H., Fujio, T., Posfai, G., Plunkett, G., Feher, T., Frisch, D., Keil, G.M., Umenhoffer, K., Kolisnychenko, V., Stahl, B., Sharma, S.S., de Arruda, M., Burland, V., Harcum, S.W. and Blattner, F.R., Emergent properties of reduced-genome Escherichia coli. Science, 2006, 312, 1044–1046.Morimoto, T., Kadoya, R., Endo, K., Tohata, M., Sawada, K., Liu, S., Ozawa, T., Kodama, T., Kakeshita, H., Kageyama, Y., Manabe, K., Kanaya, S., Ara, K., Ozaki, K. and Ogasawara, N., Enhanced recombinant protein productivity by genome reduction in Bacillus subtilis. DNA Res., 2008, 15, 73–81.

3 Kinetic Resolutions Using Biotransformations

3.1

Stereo- and Enantio-selective Hydrolysis of rac-2-Octylsulfate Using Whole Resting Cells of Pseudomonas spp. Petra Gadler and Kurt Faber

Sulfatases are a heterogenic group of hydrolytic enzymes which catalyse the cleavage of the sulfate ester bond yielding the corresponding alcohol and hydrogen sulfate. In contrast to the more commonly employed hydrolytic enzymes, such as proteases, esterases and lipases, they show not only enantioselectivity – by preference of a given substrate enantiomer over its mirror-image counterpart – but also stereoselectivity with respect to the stereochemical course of their action. Depending on the enzyme, sulfate ester hydrolysis may proceed either through retention or inversion of configuration at the chiral carbon centre (Scheme 3.1). Whereas breakage of the SO bond leads to retention, CO bond cleavage results in inversion of configuration (Scheme 3.1).1 The rare feature of double selectivities makes them top candidates for the deracemization6 of sec-alcohols via enantio-convergent hydrolysis of their corresponding sulfate esters.2 Overall, retaining sec-alkylsulfatase activity has been detected in Planctomycetes spp. (such as Rhodopirellula baltica DSM 10527;3 complementary inverting sulfatase activity was found in Actinomycetes (e.g. Rhodococcus ruber DSM 445412,4), Archaea (e.g. Sulfolobus spp.5) and pseudomonads.7 Among the latter group, Pseudomonas sp. DSM 6611 was identified as top candidate by displaying excellent stereo- and enantio-selectivities for a range of sec-alkyl sulfate esters by transforming the (R)-enantiomer of the rac-sulfate ester into the corresponding (S)-alcohol (Scheme 3.2).7



118

Kinetic Resolutions Using Biotransformations

HO 1

R

SN at Carbon Breakage of C-O bond

H 2

R

[OH– ]

H R1

Inversion

O

SO3–

R2

SN at Sulfur Breakage of S-O bond Retention

H

OH

1

R2

R

+ HSO4–

+ HSO4– –

[OH ]

Scheme 3.1 Enzymatic hydrolysis of alkylsulfate esters catalysed by alkylsulfatases proceeding through retention or inversion of configuration OSO3–

OH

Pseudomonas spp. OSO3–

E >200 Buffer pH 7.5

rac -2-octylsulfate

(S)-2-octanol OSO3–

(S)-2-octylsulfate

Scheme 3.2 Enantioselective microbial hydrolysis of rac-2-octyl sulfate using whole resting cells of Pseudomonas spp. through inversion of configuration

3.1.1 3.1.1.1

Procedure 1: Biocatalyst Preparation Materials and Equipment

• Pseudomonas spp. DSM 6611 and 6978 and Rhodococcus ruber DSM 44541 were obtained from DSMZ (Deutsche Stammsammlung fu¨r Mikroorganismen und Zellkulturen, Braunschweig, Germany, www.dsmz.de) • YPG medium comprising: – yeast extract (10 g L1) – bacteriological peptone (10 g L1) – glucose (10 g L1) – MgSO42H2O (0.15 g L1) – NaCl (2 g L1) – K2PO4 (4.4 g L1) – Na2HPO4 (1.3 g L1) • phosphate buffer (50 mM, pH 7.5; 7.58 g L1 Na2HPO42H2O and 1.01 g L1 KH2PO4) • agar plates • freeze drier. 3.1.1.2

Procedure

1. Pseudomonas spp. DSM 6611 and 6978 and Rhodococcus ruber DSM 44541 were cultivated in shaking flasks for 3 days at 30 C with shaking at 120 rpm in YPG medium containing 10 g yeast extract, 10 g bacteriological peptone, 10 g glucose,

3.1 Hydrolysis of rac-2-Octylsulfate Using Pseudomonas

119

0.15 g MgSO42H2O, 2 g NaCl, 4.4 g K2HPO4 and 1.3 g Na2HPO4 per litre. Precultures were grown either on agar plates or in liquid medium for 2–3 days as described above. 2. Cells were harvested by centrifugation for 20 min at 4 C and 8000 rpm, washed with 50 mM pH 7.5 phosphate buffer and lyophilized. Lyophilized cells were stored at 4 C. 3.1.2

Procedure 2: Microbial Hydrolysis of rac-2-Octylsulfate

3.1.2.1

Materials and Equipment

• Lyophilized whole cells of Pseudomonas spp. DSM 6611, DSM 6978 or Rhodococcus ruber DSM 44541 (50 mg) • tris-HCl buffer (600 mL, pH 7.5, 100 mM) • stock solution of substrate rac-2-octylsulfate4 (50 mg mL1 in 100 mM tris-HCl buffer pH 7.5) • ethyl acetate (600 mL) • stock solution of internal standard (10 mg mL1 rac-2-dodecanol) • Na2SO4 anhydrous. • acetic anhydride (60 mL) • 4-dimethylaminopyridine (DMAP) (cat.) • Eppendorf vials (1.5 mL) • thermoshaker • gas chromatograph (GC) equipped with a flame ionization detector (FID). 3.1.2.2

Analytics

Progress of the reaction was monitored using a GC equipped with a FID on an achiral CP 1301 capillary column (30 m 0.25 mm 0.25 mm film) and N2 as carrier gas. Enantiomeric purity of 2-octanol was analysed after derivatization with acetic anhydride (see below) using a CP-Chirasil Dex-CB column (25 m 0.32 mm 0.25 mm film, column B) and H2 as carrier gas. Enantioselectivities (expressed as the enantiomeric ratio E) were calculated from enantiomeric excess of the product and conversion as previously reported.8 Retention times and methods are listed in Table 3.1.

Table 3.1

GC methods and retention times of 2-octanol

Column

CP1301a DEX-CBb

Retention time (min) rac-2Octanol

(S)-2Octanol

(R)-2Octanol

rac-2Dodecanol

(S)-2Dodecanol

(R)-2Dodecanol

4.4 —

— 10.8

— 12.8

7.2 —

— 18.6

— 18.9

14.5 psi N2 100 C/hold 3 min – 50 C min1 – 240 C/hold 3 min. 14.5 psi H2 60 C/hold 7 min, 4 C min1 – 80 C, 10 C min1 – 160 C, 10 C min1 – 170 C/hold 5 min.

a

b

120


Table 3.2 Conversion and enantioselectivities (E-values) for the microbial hydrolysis of rac-2-octylsulfate Strain (whole cells) Pseudomonas sp. DSM 6611 Pseudomonas sp. DSM 6978 Rhodococcus ruber DSM 44541

3.1.2.3

Time (h)

Conversion (%)

Ee (S)-2-octanol (%)

24 24 24

21 7 23

99 99 77

E-value >200 >200 10

Procedure

1. Lyophilized whole cells (50 mg) of Pseudomonas spp. DSM 6611, DSM 6978 or Rhodococcus ruber DSM 44541 were rehydrated in tris-HCl buffer (600 mL, pH 7.5, 100 mM) for 1 h at 30 C with shaking at 120 rpm. 2. An aliquot (200 mL) from a substrate stock solution (50 mg mL1) was added. The mixture was incubated at 30 C with shaking at 120 rpm for 24 h. 3. The samples were extracted with ethyl acetate (600 mL) and centrifuged at 13 000 rpm for 2 min to separate the organic layer from the cell/buffer suspension. The organic layer was dried over Na2SO4 and 100 mL of an internal standard (10 mg mL1 rac-2dodecanol) was added. 4. Conversions were measured on an achiral GC column using calibration curves. For the determination of the enantiomeric excess, the 2-octanol formed was derivatized into the corresponding acetate ester using acetic anhydride (60 mL) and catalytic DMAP overnight. The reaction was quenched with tap water (300 mL), centrifuged for 2 min at 13 000 rpm and the organic layer dried (Na2SO4) and analysed as described above. Results are listed in Table 3.2.

References 1. Gadler, P. and Faber, K., New enzymes for biotransformations: microbial alkyl sulfatases displaying stereo- and enantioselectivity. Trends Biotechnol., 2007, 25, 83. 2. Wallner, S.R., Pogorevc, M., Trauthwein, H. and Faber, K., Biocatalytic enantioconvergent preparation of sec-alcohols using sulfatases. Eng. Life Sci., 2004, 4, 512. 3. Wallner, S.R., Bauer, M., Wu¨rdemann, C., Wecker, P., Gloeckner, F.O. and Faber, K., Highly enantioselective sec-alkyl sulfatase activity of the marine planctomycete Rhodopirellula baltica shows retention of configuration. Angew. Chem. Int. Ed., 2005, 44, 6381. 4. Pogorevc, M. and Faber, K., Enantioselective stereoinversion of sec-alkyl sulfates by an alkylsulfatase from Rhodococcus ruber DSM 44541. Tetrahedron Asymm., 2002, 13, 1435. 5. (a) Wallner, S.R., Nestl, B.M. and Faber, K., Highly enantioselective sec-alkyl sulfatase activity of Sulfolobus acidocaldarius DSM 639. Org. Lett., 2004, 6, 5009; (b) Wallner, S.R., Nestl, B.M. and Faber, K., Highly enantioselective stereo-inverting sec-alkylsulfatase activity of hyperthermophilic Archaea. Org. Biomol. Chem., 2005, 3, 2652. 6. (a) Faber, K., Non-sequential processes for the transformation of a racemate into a single stereoisomeric product: proposal for stereochemical classification. Chem. Eur. J., 2001, 7, 5004; (b) Gadler, P., Glueck, S.M., Kroutil, W., Nestl, B.M., Larissegger-Schnell,B., Ueberbacher, B.T., Wallner, S.R. and Faber, K., Biocatalytic approaches for the quantitative production of single stereoisomers from racemates. Biochem. Soc. Trans., 2006, 34, 296. 7. Gadler, P. and Faber, K., Highly enantioselective biohydrolysis of sec-alkyl sulfate esters with inversion of configuration catalysed by Pseudomonas spp. Eur. J. Org. Chem., 2007, 5527. 8. Chen, C.-S., Fujimoto, Y., Girdaukas, G. and Sih, C.J., Quantitative analysis of biochemical kinetic resolutions of enantiomers. J. Am. Chem. Soc., 1982, 104, 7294.

3.2 Protease-catalyzed Resolutions Using p-Toluenesulfonamide

3.2

121

Protease-catalyzed Resolutions Using the 3-(3-Pyridine)propionyl Anchor Group: p-Toluenesulfonamide Christopher K. Savile and Romas J. Kazlauskas

Proteases require water-soluble substrates and bind them in a shallow active site in an extended conformation.1 The shallow active site allows proteases to accept sterically hindered substrates2,3 and also polar substrates, since one substituent remains in water.4 To bind substrates, proteases contain a specificity pocket for the acyl group.5,6 For example, subtilisins and chymotrypsin favour ester and amides of phenylalanine.5–7 The 3-(3-pyridine)propionyl group mimics phenylalanine and increases substrate binding and solubility in water, thereby increasing the rates of protease-catalyzed reactions. In addition, the 3-(3-pyridine)propionyl group eliminates chromatography, since mild acid extraction separates the remaining starting material and product. To demonstrate the synthetic usefulness of this strategy, we resolved multi-gram quantities of (R)- and (S)-p-toluenesulfinamide with -chymotrypsin.8 3.2.1

Procedure: Resolution of N-3-(3-Pyridine)propionyl-p-tolylsulfinamide O

OH

carbodiimide

O

O O

N

N

N O

O H 3C

O S N H

1

S NH 2

20.2g 1a

NaH

α-chymotrypsin H2O E=52

H3 C

3.81 g 35 % yield 98 % ee af ter H 3C recrystallization

S (S)

NH2 (S)-1 3.58 g 33 % yield 98 % ee af ter recrystallization

3.2.1.1 • • • • •

(R)

O

S NH 2

(R)-1 + O

O H3 C

N

O S N H

(1) H Extraction (2) NH2NH2

N H3 C


N-(3-Dimethylaminopropyl)-N0 -ethylcarbodiimide hydrochloride (25.2 g, 132 mmol) triethylamine (37 mL) 3-(3-pyridine)propionic acid (40.0 g, 265 mmol) CH2Cl2 (3250 mL) p-toluenesulfinamide (15.5 g, 100 mmol)

122

• • • • • • • • • • • • • • • • • • • • • • • • • • • • • • • •


sodium hydride (60 % dispersion in oil; 12.0 g, 300 mmol) tetrahydrofuran (THF, 750 mL) EtOAc (1550 mL) hexanes (150 mL) aqueous saturated NaHCO3 (1400 mL) MgSO4 anhydrous bovine -chymotrypsin, Sigma (12 g) N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES, 0.7 g) potassium chloride (23.4 g) dimethylformamide (350 mL) NaOH, 1.0 M (35 mL) aqueous saturated NaCl (750 mL) HCl, 0.10 M (200 mL) hydrazine hydrate (35 mL) HCl, 1.0 M (50 mL) three-neck round-bottom flask, 250 mL and 2 L with magnetic stirbar glass stopper (24/40) rubber septum (24/40) cannula (18 gauge) syringe and needle (18 gauge) ice bath magnetic stir plate graduated cylinder, 1 L separatory funnel, 500 mL and 2 L Erlenmeyer flask, 1 L and 2 L glass funnel filter paper round-bottom flask, 1 L and 2 L rotary evaporator beaker, 4 L with magnetic stir bar dropping funnel, 500 mL pH Stat (Radiometer Titralab TIM854 or equivalent).

3.2.1.2

Procedure

3-(3-Pyridine)propionic Acid Anhydride. 1. N-(3-Dimethylaminopropyl)-N0 -ethylcarbodiimide hydrochloride (25.2 g, 132 mmol) was added to a solution of 3-(3-pyridine)propionic acid (40.0 g, 265 mmol) in CH2Cl2 (750 mL) and Et3N (37 mL) at 0 C and stirred at room temperature (RT).8,9 2. After 24 h, the reaction was washed with ice-cold saturated NaHCO3 (3 500 mL), dried over MgSO4 and concentrated in vacuo to give a pale yellow oil (34.1 g, 91 %). 1 H NMR 2.72 (t, J ¼ 7.2, 2H, C(O)CH2), 3.03 (t, J ¼ 7.5, 2H, CH2Ph), 7.24 (m, 1H, pyridyl), 7.58 (m, 1H, pyridyl), 8.50 (m, 2H, pyridyl).

3.2 Protease-catalyzed Resolutions Using p-Toluenesulfonamide

123

Racemic N-3-(3-Pyridine)propionyl-p-toluene sulfinamide 1a. 1. Sodium hydride (60 % dispersion in oil; 12.0 g, 300 mmol) was added portion-wise over 15 min to a solution of p-toluenesulfinamide (15.5 g, 100 mmol) in THF (750 mL) at 0 C. The symmetrical anhydride of 3-(3-pyridine)propionic acid (32.1 g, 113 mmol) was added drop-wise over 15 min at 0 C and the reaction mixture was then stirred at RT for 3 h.10 2. The reaction mixture was diluted with EtOAc (400 mL) and saturated NaHCO3 (400 mL) was added slowly. The layers were separated and the aqueous layer was extracted with EtOAc (3 250 mL). The combined EtOAc layers were washed with saturated NaHCO3 (500 mL) and dried over MgSO4. The aqueous layer was extracted with CH2Cl2 (2 250 mL). The combined CH2Cl2 layers were washed with NaHCO3 (250 mL) and dried over MgSO4. The combined organic layers were concentrated in vacuo to give a pale yellow solid. Trituration with hexane/ethyl acetate gave 1a as a white powder (21.1 g, 73 %). Mpt. 161–163 C; 1H NMR 2.39 (s, 3H, PhCH3), 2.72 (m, 2H, C(O)CH2), 3.01 (t, J ¼ 7.2, 2H, CH2Pyr), 4.78 (br s, 1 H, NH), 7.25–7.48 (m, 3H, phenyl or pyridyl), 7.47 (m, 2H, phenyl or pyridyl), 7.68 (m, 1H, phenyl or pyridyl), 8.25 (m, 2H, pyridyl); 13 C NMR (DMSO-d6) 21.4 (PhCH3), 27.9 (CH2Pyr), 36.9 (C(O)CH2), 124.0, 125.4, 130.2, 136.4, 136.6, 140.9, 142.1, 147.9, 150.2 (phenyl or pyridyl), 173.8 (C¼O); HRMS calc. for C15H17N2O2S [M þ H]þ 289.1010. Found: 289.0989. The enantiomers were separated using HPLC (Chiralcel OD-H column, 85:15 hexanes/EtOH, 0.75 mL min1, 254 nm; (R)-enantiomer tR ¼ 20.0 min; (S)-enantiomer, tR ¼ 22.5 min). Resolution of N-3-(3-Pyridine)propionyl-p-tolylsulfinamide. 1. -Chymotrypsin (12 g) was added to a solution of BES buffer (3.15 L, 1 mM, pH 7.2) and 100 mM KCl and stirred for 15 min to ensure complete dissolution. Substrate 1a (20.2 g, 70 mmol) was dissolved in dimethylformamide (350 mL) and added drop-wise to the enzyme solution. The rate of hydrolysis was monitored by pH Stat, which maintained the pH at 7.2 by automatic titration with 1 M NaOH. (i) Subtilisin BPN0 or subtilisin E are better proteases for this resolution because they are more enantioselective, but they require a fermentation to produce.11 As an alternative, we chose a commercially available, but less enantioselective, protease – -chymotrypsin. (ii) If a pH Stat is not available, increase the buffer concentration from 1 mM to 50 mM and maintain the pH at 7.2 by manual addition of 1 M NaOH. Approximately 35 mL will be required. 2. At 50 % conversion (4 days), the reaction was terminated by extraction of remaining starting material and product with CH2Cl2 (3 500 mL). The combined organic layers were washed with H2O (3 500 mL), saturated NaCl (1 500 mL), dried over MgSO4 and concentrated in vacuo. The crude mixture was dissolved in EtOAc (250 mL) and unreacted starting material was extracted with ice-cold 0.1 M HCl (2 100 mL). The combined aqueous layers were then back-extracted with EtOAc (50 mL). The combined EtOAc layers were washed with saturated NaHCO3 (100 mL), saturated NaCl (100 mL), dried over MgSO4 and concentrated in vacuo to give (R)-1 as a white solid (4.48 g, 41 % yield) with 87 % ee.

124


3. The combined aqueous layers were neutralized with solid NaHCO3 and extracted with CH2Cl2 (2 200 mL). The combined CH2Cl2 layers were washed with saturated NaHCO3 (100 mL), saturated NaCl (100 mL) and dried over MgSO4. The solution was concentrated in vacuo to give (S)-1a, which was subsequently treated with hydrazine hydrate (35 mL).12 After stirring for 3 h, the reaction solution was diluted with CH2Cl2 (100 mL) and washed with 1 M HCl (50 mL), saturated NaHCO3 (50 mL), saturated NaCl (50 mL) and concentrated in vacuo to give (S)-1 (4.29 g, 40 % yield) with 92 % ee. 4. Recrystallization from hexanes/ethyl acetate gave (R)-1 (3.81 g, 35 % yield) with 98% ee and (S)-1 (3.58 g, 33 % yield) with 98 % ee.

References 1. Tyndall, J.D.A., TessaNall, T. and Fairlie, D.P., Proteases universally recognize beta strands in their active sites. Chem. Rev., 2005, 105, 973–1000. 2. (a) Muchmore, D.C., Enantiomeric enrichment of (R,S)-3-quinuclidinol. US Patent US 5,215,918, 1993; (b) Savile, C.K., Magloire, V.P. and Kazlauskas, R.J., Subtilisin-catalyzed resolution of N-acyl arylsulfinamides. J. Am. Chem. Soc., 2005, 127, 2104–2113. 3. Mugford, P.F., Lait, S.M., Keay, B.A. and Kazlauskas, R.J., Enantiocomplementary enzymatic resolution of the chiral auxiliary cis,cis-6-(2,2-dimethylpropanamido)spiro-[4.4]nonan-1-ol and the moleuclar basis for the high enantioselectivity of subtilisin Carlsberg. ChemBioChem, 2004, 5, 980–987. 4. Savile, C.K. and Kazlauskas, R.J., How substrate solvation contributes to the enantioselectivity of subtilisin toward secondary alcohols. J. Am. Chem. Soc., 2005, 127, 12228–12229. 5. (a) Lin, Y.Y., Palmer, D.N. and Jones, J.B., The specificity of the nucleophilic site of -chymotrypsin and its potential for the resolution of alcohols. Enzyme-catalyzed hydrolyses of some (þ)-, ()-, and (–)-2-butyl, -2-octyl, and --phenethyl esters. Can. J. Chem., 1974, 52, 469–476. 6. (a) Estell, D.A., Graycar, T.P., Miller, J.V., Powers, D.B., Burnier, J.P., Ng, P.G. and Wells, J.A., Probing steric and hydrophobic effects on enzyme–substrate interactions by protein engineering. Science, 1986, 233, 659–663. (b). Wells, J.A., Powers, D.B., Bott, R.R., Graycar, T.P. and Estell, D.A., Proc. Natl. Acad. Sci. U. S. A., Designing substrate specificity by protein engineering of electrostatic interactions. 1987, 84, 1219–1223. 7. Pohl, T. and Waldmann, H., Enhancement of the enantioselectivity of penicillin G acylase from E. coli by ‘substrate tuning’. Tetrahedron Lett., 1995, 36, 2963–2966. 8. Savile, C. K., Kazlauskas, R.J., The 3-(3-pyridine)propionyl anchor group for protease-catalyzed resolutions: p-toluenesulfinamide and sterically hindered secondary alcohols. Adv. Synth. Catal., 2006, 348, 1183–1192. 9. Walker, F.A. and Benson, M., Entropy, enthalpy, and side arm porphyrins. 1. Thermodynamics of axial ligand competition between 3-picoline and a series of 3-pyridyl ligands covalently attached to zinc tetraphenylporphyrin. J. Am. Chem. Soc., 1980, 102, 5530–5538. 10. Backes, B.J., Dragoli, D.R. and Ellman, J.A., Chiral N-acyl-tert-butanesulfinamides: the ‘safety-catch’ principle applied to diastereoselective enolate alkylations. J. Org. Chem., 1999, 64, 5472–5478. 11. (a) Harwood, C.R. and Cutting, S.M., Molecular Biological Methods for Bacillus, John Wiley & Sons, Ltd, Chichester, 1990, pp. 33–35, 391–402; (b) Cho, S.-J., Oh, S.-H., Pridmore, R.D., Juillerat, M.A. and Lee,C.[hyphen]H., Purification and characterization of proteases from Bacillus amyloliquefaciens isolated from traditional soybean fermentation starter. Agric. Food Chem., 2003, 51, 7664–7670. 12. Keith, D.D., Tortora, J.A. and Yang, R., Synthesis of L-2-amino-4-methoxy-trans-but-3-enoic acid. J. Org. Chem., 1978, 43, 3711–3713.

3.3 Desymmetrization of Prochiral Ketones Using Enzymes

3.3

125

Desymmetrization of Prochiral Ketones Using Enzymes Andrew J. Carnell

Chiral enol esters are useful synthetic intermediates which can serve as chiral enolate equivalents or undergo oxidative cleavage to produce reactive ester-aldehydes.1,2 Enolate equivalents, such as silyl enol ethers and enol esters, can be made using chiral lithium amide bases at low temperature, although poor selectivity can be a problem, particularly with nonconformationally locked ketones.3 Lipase-catalysed resolution by enantioselective transesterification of racemic enol acetates derived from prochiral ketones can give enol acetates in very high ee.4 Enol esters derived from 8-oxabicyclic ketones can be resolved in excellent selectivity (E ¼ 45–48) using silica-absorbed butanol-rinsed enzyme preparation (BREP) Humicola sp. lipase.5 Similarly, enol esters derived from 4,4-disubstituted cyclohexanones can be resolved using Pseudomonas fluorescens lipase (PFL) in tetrahydrofuran (THF).6,7 The prochiral ketone can be recycled, leading to a formal desymmetrization of the ketone and good yield of the enantiomerically pure enol ester. O

R

R

O

+

OAc AcO

R

R

O

Humicola sp. lipase (BREP)

Ar CN (S)

3.3.1.1 • • • • • • •

THF E = 6.5–13

(R)

O R1

Ar CN

N

steps

Ar CN (S)

recycle

R 4 R = Me, 5 R = Et 6 R = CH2OMe

OAc +

Ar CN

O

(1R, 5S)

PFL, n-BuOH

7 Ar = Ph 8 Ar = 3,4-Cl2C6H 3 9 Ar = 3,4-(MeO)2C6H 3

3.3.1

O

O

+

91-99% e.e.

OAc +

R OAc

R

hexane/n-BuOH E = 45-48

1 R = Me, 2 R = Et 3 R = CH2OMe

OAc

R

NK-2 antagonists

Ar NR2

95–99% e.e. 10 Ar = Ph 11 Ar = 3,4-Cl2C6H 3 (82%, 99% e.e. after 3 recycles of ketone) 12 Ar = 3,4-(MeO)2C6H 3

Procedure 1: Humicola sp. Lipase BREP Materials and Equipment

Humicola sp. lipase (Chirazyme L-8) (190 mg) silica gel (Merck Kieselgel 60 (230–400 mesh) (3.2 g) tris-HCl buffer (pH 7, 64 mL) dry n-butanol (120 mL) dry n-hexane (120 mL) one 250 mL conical flask orbital shaker.

3.3.1.2

Procedure

1. Humicola sp. lipase (190 mg) was dissolved in tris-HCl buffer (pH 7, 64 mL). To this solution was added silica gel (3.2 g) and the mixture shaken at room temperature for 2 h.

126


2. The mixture was allowed to settle and the aqueous phase decanted ensuring that the silica remained wet. 3. Dry n-butanol (40 mL) was added, swirled and decanted ensuring that the silica remained wet. This was repeated twice (2 40 mL of n-butanol). 4. Dry hexane (40 mL) was added, swirled and decanted ensuring that the silica remained wet. This was repeated twice (2 40 mL of hexane) 3.3.2 3.3.2.1 • • • • • • • • • • • •

Procedure 2: Resolution of (–)-1,5-Dimethyl-3-acetyloxy-8oxabicyclo[3.2.1]oct-2,6-diene (1) using Humicola sp. Lipase BREP Materials and Equipment

One quantity of Humicola sp. lipase BREP prepared as above (–)-1,5-dimethyl-3-acetyloxy-8-oxabicyclo[3.2.1]oct-2,6-diene 1 (760 mg, 3.92 mmol) dry n-butanol (0.72 mL, 7.84 mmol) dry n-hexane (40 mL) Celite (5 g) hexane for washing (30 mL) two 100 mL round-bottom flasks stirrer bar magnetic stirrer plate sintered vacuum filtration funnel rotary evaporator equipment for column chromatography.

3.3.2.2

Procedure

1. Dry hexane (40 mL) was added to the Humicola sp. lipase BREP prepared above in Procedure 1. This suspension was transferred to a 100 mL round-bottom flask containing the enol acetate 1 (760 mg, 3.92 mmol). Dry n-butanol (0.72 mL, 7.84 mmol) was added and the mixture stirred at 25 C for 1 h. 2. The mixture was filtered through a pad of Celite (5 g) using a vacuum sinter funnel and washed through into a 100 mL flask with dry hexane (3 30 mL). The filtrate was concentrated in vacuo to yield a crude mixture which was separated using flash column chromatography on silica with 5 % ethyl acetate/40–60 % petroleum ether as eluent to give the prochiral ketone 4 (290 mg, 49%) and enol acetate ()-1 of >99.5 % ee (230 mg, 30 %) as a yellow oil []D 53.5 (c ¼ 8, CHCl3); (high-resolution mass spectrometry (HRMS): Found [M þ H]þ 195.10237. C11H15O3 requires 195.10212); max (neat)/cm1 1759 (CO); H (300 MHz; C6D6; Me4Si) 1.29 (3 H, s, CH3), 1.33 (3 H, s, CH3), 1.66 (3 H, s, COCH3), 1.83 (1 H, dd, J 17 and 1.5, CH2endo), 2.45 (1 H, dd, J 17 and 1.8, CH2exo), 5.43 (1 H, d, J 5.7, CH¼), 5.81 (1 H, d overlapping, J 1.8 and 1.5, CH¼CO) and 5.99 (1 H, d, J 5.4, CH¼); C (75.5 MHz; C6D6; Me4Si) 20.8 (CH3), 22.0 (CH3), 24.4 (CH3), 36.8 (CH2), 82.1 (C), 83.3 (C), 121.9 (CH), 131.6 (CH), 141.4 (CH), 147.7 (C) and 168.6 (C); m/z (CI) 212 (100 %, [M þ NH4]þ), 195 (37, [M þ H]þ). Chiral analysis was done using a Chiralpak AD column eluting with 5–10 % isopropanol/n-hexane (1 mL min1) and was recorded at 220 nm.

3.3 Desymmetrization of Prochiral Ketones Using Enzymes

3.3.3 3.3.3.1 • • • • • • • • • • •

127

Procedure 3: Resolution of (–)-4-Cyano-4-(30 ,40 -dichlorophenyl)cyclohex-1enyl Acetate (8) Materials and Equipment

(–)-4-Cyano-4-(30 ,40 -dichlorophenyl)cyclohex-1-enyl acetate (8) (10 g, 32.4 mmol) PFL (Amano AK) (8 g) n-butanol (5.21 mL, 64.7 mmol) THF (125 mL) THF for washing (100 mL) 250 mL round-bottom flask stirrer bar magnetic stirrer plate sintered vacuum filtration funnel rotary evaporator equipment for column chromatography.

3.3.3.2

Procedure

1. The (–)-enol acetate 8 (1 g, 32.4 mmol), PFL (8 g) and n-BuOH (5.21 mL, 64.7 mmol) were stirred in THF (125 mL) at room temperature for 9.5 h. 2. The solution was filtered through a glass sinter funnel under vacuum, the residual enzyme washed with THF (100 mL) and the solvent removed by evaporation under reduced pressure. The crude residue was purified by flash chromatography on silica using diethyl ether/petroleum ether (1:2) as eluent to give the ketone (7 g) and the (S)-enol acetate 8 (2.8 g, 28 %, >99 % ee) as a white solid, m.p. 148–150 C, []D ¼ þ11.5 (c ¼ 1.74 in CHCl3). Chiral HPLC (Chiralpak AD) indicated 100 % ee for the enol acetate. RT (R)-8 15.5 min. (S)-8 20.9 min, eluent 100% EtOH, flow rate 0.5 mL min1, l 220 nm. (Found: C, 57.81; H, 4.18; N, 4.48. C15H13Cl2NO2 requires C, 58.08; H, 4.22; N, 4.52 %) (HRMS: found Mþ 309.0322. C15H1335Cl2NO2 requires 309.0323); max(neat) 2233 (CN), 1755 (CO); H (300 MHz, CDCl3) 2.23 (3 H, s, AcCH3), 2.23–2.67 (6 H, m, 2 H-3, 2 H-5 and 2 H-6), 5.46 (1H, s, H-2) 7.22–7.56 (3 H, Ar); C (75 MHz, CDCl3) 20.9 (AcCH3), 24.6, 32.7, 35.4 (C-3, C-5, and C-6), 40.0 (C-4), 110.5 (C-2), 121.5 (CN), 125.3, 128.1 and 131.1 (C-20 , C-50 , C-60 ), 132.8, 133.5 and 139.6 (C-10 , C-30 and C-40 ), 148.1 (C-1), 169.3 (C¼O); m/z (EI) 309 (3 %, Mþ), 267 (11), 70 (69), 43 (100). 3.3.4

Conclusion

This methodology was shown to work well for the desymmetrization of related ketones. For example, oxabicyclic enol acetates 2 and 3 with other substituents (Et and CH2OMe) at the bridge position were transformed more slowly but with similarly high enantioselectivity (Table 3.3).3 For the cyclohexanone series, other aryl groups are tolerated at the 4-position as in substrates 7 and 9, although with PFL the cyano group is required for good enantioselectivity (Table 3.4).1 An attractive feature of this type of resolution is that the prochiral ketone can be recycled. The homochiral (S)-enol ester 8 was obtained in 82 % yield by recycling the ketone without prior separation from the enantioenriched enol ester. For a cyclic enzyme

128


Table 3.3 Reactions carried out in hexane using Humicola sp. lipase as described in Procedure 2 Substrate 1 2 3

Reaction time (h)

Conversion (%)

ee of enol acetate (%)

E

1 19 48

67 51 53

>99 91 96

45 47 48

Table 3.4 Reactions carried out in THF using freeze-dried Amano AK PFL as described in Procedure 3 Substrate 7 8 9

Conversion to ketone (%)

ee of enol acetate (%)

E

68 70 71

>99 (S) >99 (S) 95

13 11 7.4

resolution of this type it can be derived that the maximum theoretical enantiomeric excess for 100 % yield is eemax ¼ (E 1)/(E þ 1).6 Thus, for an enzyme resolution with an E value of 13, the eemax ¼ 85.7 %. A higher ee in a stepwise process is possible only if the yield is compromised. This ee corresponds to a conversion of 56 % and is the optimum point at which to stop each kinetic resolution. Of course, with a mixture of ketone and enantioenriched ester for the start of each biotransformation after the first, the conversion required to get to the 56 % (or 85.7 % ee) point is less. In the final biotransformation the conversion is allowed to go beyond this point and the yield is compromised in order to get homochiral ester. The enol ester 8 was subsequently used in a short and efficient four-step synthesis of a nonpeptidic neurokinin NK-2 antagonist developed by Pfizer for the treatment of neuroinflammatory conditions.1,2

References 1. Allan, G., Carnell, A.J, Escudero Hernandez, M.L. and Pettman, A., Chemoenzymatic synthesis of a tachykinin NK-2 antagonist. Tetrahedron, 2001, 57, 8193. 2. Carnell, A.J., Escudero Hernandez, M.L., Pettman, A. and Bickley, J., Chemoenzymatic synthesis of a non-peptide tachykinin NK-2 antagonist. Tetrahedron Lett., 2000, 41, 6929. 3. Allan, G., Carnell, A.J., Escudero Hernandez, M.L. and Pettman, A., Desymmetrisation of 4,4disubstituted cyclohexanones by enzyme-catalysed resolution of their enol acetates. J. Chem. Soc. Perkin Trans. 1, 2000, 3382. 4. Carnell, A., Desymmetrisation of prochiral ketones using lipases. J. Mol. Catal. B Enzymatic, 2002, 19–20, 83. 5. Carnell, A.J., Swain, S.A. and Bickley, J.F., Chiral enol acetates derived from prochiral oxabicyclic ketones using enzymes. Tetrahedron Lett., 1999, 40, 8633. 6. Carnell, A.J., Barkley, J. and Singh, A., Desymmetrisation of prochiral ketones by catalytic enantioselective hydrolysis of their enol esters using enzymes. Tetrahedron Lett., 1997, 38, 7781. 7. Allan, G., Carnell, A.J. and Kroutil, W., One-pot deracemisation of an enol acetate derived from a prochiral cyclohexanone. Tetrahedron Lett., 2001, 42, 5959.

3.4 Enzymatic Resolution of 1-MTQ using C rugosa Lipase

3.4

129

Enzymatic Resolution of 1-Methyl-tetrahydroisoquinoline using Candida rugosa Lipase Gary Breen

Although secondary amines are common building blocks in the pharmaceutical industry, there are few examples of the resolution of secondary amines in the literature. Preparation of substituted phenyl allylcarbonates allowed the resolution of 1-methyl-tetrahydroisoquinoline (1-MTQ) to proceed with excellent enantioselectivity and recovery (Figure 3.1). 3.4.1

Procedure 1: Preparation of 3-Methoxyphenyl Allylcarbonate1 O MeO

3.4.1.1 • • • • • • • • • • • • • •

O

O


3-Methoxyphenol (6.21 g) tetra-n-butylammonium chloride hydrate (100 mg) dichloromethane (40 mL) allyl chloroformate (6 mL) 4 M sodium hydroxide solution (30 mL) anhydrous magnesium sulfate N2 gas one 100 mL three-necked flask with a magnetic stirrer one magnetic stirring hotplate ice one 100 mL separating funnel filter paper rotary evaporator Kugelrohr distillation equipment.

3.4.1.2

Procedure

1. 3-Methoxyphenol (6.21 g) and tetra-n-butylammonium chloride hydrate (100 mg) were dissolved in dichloromethane (40 mL) in a 100 mL three-necked flask. 2. Sodium hydroxide solution (4 M, 20 mL) was added and the mixture cooled to 0–5 C in an ice bath with magnetic stirring under nitrogen. 3. Allyl chloroformate (6 mL) was added slowly, keeping the temperature between 0 and 5 C. 4. After stirring for a further 1 h, the two layers were separated in a 100 mL separating funnel and the dichloromethane layer was washed with 10 mL 4 M sodium hydroxide solution. The organic layer was then dried with anhydrous magnesium sulfate and concentrated using a rotary evaporator. The crude product (10.0 g) was purified using vacuum distillation on a Kugelrohr apparatus (3 mbar, 100 C, cooling the recipient flask with ice). (Yield 9.2 g, 88 %.) 1 H NMR (400 MHz, CDCl3) 3.80 (3H, s), 4.75 (2H, d, J 8.3 Hz), 5.31–5.44 (2H, m), 5.95–6.05 (1H, m), 6.74 (1H, t, J 4.8 Hz), 6.77–6.81 (2H, m), 7.28 (1H, d, J 8.8 Hz).

130

Kinetic Resolutions Using Biotransformations O R

O

O NH

NH

+

Candida rugosa lipase

O O

(S)-1-MTQ

1-MTQ

Figure 3.1

3.4.2

N

Enzymatic resolution of 1-MTQ

Procedure 2: Synthesis of (S)-1-Methyltetrahydroisoquinoline

NH (S)-1-MTQ

3.4.2.1 • • • • • • • • • • • • • • •


Racemic 1-methyltetrahydroisoquinoline (5 g) toluene (water saturated, 70 mL)2 3-methoxyphenyl allylcarbonate (4.65 g) ChiroCLEC-CR (100 mg)3 saturated sodium chloride solution (50 mL) 2 M hydrochloric acid solution (50 mL) 10 M sodium hydroxide solution tert-butylmethylether (TBME, 100 mL) anhydrous magnesium sulfate two 100 mL round-bottomed flasks two magnetic stirring hotplates one Bu¨chner flask, 100 mL one Bu¨chner funnel one 100 mL separating funnel rotary evaporator.

3.4.2.2

Procedure

1. Racemic 1-methyltetrahydroisoquinoline (5 g) and 3-methoxyphenyl allylcarbonate (4.65 g) were stirred at 30 C in a round-bottomed flask connected in a closed system to another round-bottomed flask containing saturated sodium chloride solution at 50 C.4 2. ChiroCLEC-CR was added to the reaction flask and the reaction monitored for completion by high-performance liquid chromatography (HPLC). 3. After 8 h the enzyme was filtered off in a Bu¨chner funnel and washed with toluene (10 mL). The combined organic layers were washed with 2 M hydrochloric acid solution (2 25 mL) in a 100 mL separating funnel. The combined acid layers were then washed with toluene (10 mL) and the pH then adjusted to 12 with 10 M sodium

3.4 Enzymatic Resolution of 1-MTQ using C rugosa Lipase

131

hydroxide solution. The oil which formed was extracted with TBME (2 50 mL). The organic portion was dried over anhydrous magnesium sulfate and concentrated using a rotary evaporator. The product, (S)-1-MTQ was obtained as an oil with no further purification. (Yield 2.3 g, 46 %, 99.6 % ee.) 1 H NMR (400 MHz, CDCl3) 1.46 (3H, d, J 6.8 Hz), 1.90 (1H, br s), 2.73 (1H, dt, J 16.3, 4.8 Hz, 2.87 (1H, m), 3.02 (1H, m), 3.26 (1H, dt, J 12.8, 5.0 Hz), 4.10 (1H, q, J 6.8 Hz), 7.10 (4H, m). HPLC analysis: Chiralcel OD column, 3 % hexane in methanol eluent, 1.5 mL min1, UV at 220 nm. Typical retention times: (S)-1-MTQ, 8.6 min; (R)-1-MTQ, 10.4 min. 3.4.3

Conclusion

This is a simple procedure for the enzymatic resolution of a secondary amine. The acylating agent can be modified by altering the substitution on the phenol ring. This tuning of the reactivity and selectivity should allow other amines to be resolved using a similar approach.

References and Notes 1. This acylating agent has also been used in the resolution of indolines; see Gotor-Fernandez, V., Rebolledo, F. and Gotor, V., Chemoenzymatic preparation of optically active secondary amines: a new efficient route to enantiomerically pure indolines. Tetrahedron Lett., 2006, 17, 2558. 2. This is prepared by stirring toluene with excess water and separating the two layers. The saturated toluene layer contains 0.05 % w/w water. Water is important to maintain the conformational integrity of the enzyme. 3. This enzyme is no longer commercially available, but other C. rugosa lipases were also found to be active under these conditions, including Biocalysts L034P (LipomodTM 34P). 4. The saturated salt solution maintains a constant water level in the toluene solution and leads to faster reaction times. Section 3.4 reprinted from Breen, G. F. Enzymatic resolution of a secondary amine using novel acylating reagents. Tetrahedron Asymmetry 2004, 15(9), 1427–1430, with permission from Elsevier.

4 Dynamic Kinetic Resolution for the Synthesis of Esters, Amides and Acids Using Lipases

4.1

Dynamic Kinetic Resolution of 1-Phenylethanol by Immobilized Lipase Coupled with In Situ Racemization over Zeolite Beta Kam Loon Fow, Yongzhong Zhu, Gaik Khuan Chuah and Stephan Jaenicke

The one-pot dynamic kinetic resolution (DKR) of (–)-1-phenylethanol lipase esterification in the presence of zeolite beta followed by saponification leads to (R)-1 phenylethanol in 70 % isolated yield at a multi-gram scale. The DKR consists of two parallel reactions: kinetic resolution by transesterification with an immobilized biocatalyst (lipase B from Candida antarctica) and in situ racemization over a zeolite beta (Si/Al ¼ 150).1 With vinyl octanoate as the acyl donor, the desired ester of (R)-1-phenylethanol was obtained with a yield of 80 % and an ee of 98 %. The chiral secondary alcohol can be regenerated from the ester without loss of optical purity. The advantages of this method are that it uses a single liquid phase and both catalysts are solids which can be easily removed by filtration. This makes the method suitable for scale-up. The examples given here describe the multi-gram synthesis of (R)-1-phenylethyl octanoate and the hydrolysis of the ester to obtain pure (R)-1-phenylethanol.



134

DKR for the Synthesis of Esters, Amides and Acids Using Lipases

4.1.1

Procedure 1: Synthesis of (R)-1-Phenylethyl Octanoate O OH

Catalyst: Novozym 435 Zeolite Beta

O

+

O

C7H15

O

C 7H 15 O

+ Toluene, 60 °C

H 80% yield 98% ee

4.1.1.1 • • • • • • • • • • • • • • •


(–)-1-Phenylethanol (1.22 g, 10 mmol) vinyl octanoate (2.04 g, 12 mmol) H-zeolite beta with Si/Al ¼ 150 (250 mg) immobilized lipase B from C. antarctica (E.C.3.1.1.3); tradename: Novozym 435 (150 mg) toluene (5 mL) hexane (20 mL) 0.1 M NaOH solution (60 mL) Na2SO4, anhydrous (3 g) two-necked round-bottom flask, 25 mL capacity magnetic stirring bar hotplate with magnetic stirrer oil bath analytical balance separation funnel, 50 mL rotary evaporator.

Vinyl octanoate was obtained from TCI (Tokyo, Japan). All other chemicals with the exception of the zeolite beta are available from Sigma Aldrich. The synthesis of a particularly active modification of low-alumina zeolite beta has been described by us.2 Commercial material, available as samples from, for example, Zeolyst or Su¨dchemie can be used, but because of excessive acidity may result in up to 15 % of styrene formation. 4.1.1.2

Procedure

1. (–)-1-Phenylethanol (1.22 g, 10 mmol), vinyl octanoate (2.04 g, 12 mmol) and toluene (5 mL) were added into a 50 mL round-bottom flask and heated to 60 C in an oil bath. Zeolite beta with Si/Al ¼ 150 (250 mg) and the immobilized lipase Novozym 435 (150 mg) were added to the reaction mixture. The mixture was stirred for 6 h at 60 C. 2. The mixture was left to cool to room temperature and the solid catalysts were removed by filtration. Toluene and the by-product, acetaldehyde, were removed under reduced pressure by using a rotary evaporator. The residue contains the desired product together with unreacted vinyl octanoate and traces of octanoic acid, which are formed by hydrolysis of the vinyl octanoate. The product can be purified by redissolving the residue in hexane (20 mL) and washing it with 0.1 M NaOH (20 mL 3). The organic layer was dried with anhydrous Na2SO4 and the hexane was removed by a rotary evaporator.

4.1 DKR of 1-Phenylethanol by Immobilized Lipase Coupled

135

3. The product is pure (R)-1-phenylethyl octanoate (2.01 g, 80 % yield). 1 H NMR (300 MHz, CDCl3) 7.42–7.40 (m, 5H), 5.98 (q, J ¼ 6.6, 1H), 2.39 (t, J ¼ 7.1, 2H), 1.60 (d, J ¼ 6.8, 3H), 1.36-1.35 (m, 10H), 0.96 (t, J ¼ 6.6, 3H). 13 C NMR (75 MHz, CDCl3) 173.53, 142.45, 129.00 (2 C), 128.31, 126.60 (2 C), 72.53, 35.15, 32.23, 29.61, 29.48, 25.55, 23.15, 22.80, 14.61. The purity and ee were determined by gas chromatography (GC) with a chiral column (Supelco Beta DEX 120 (90 C isotherm)): major enantiomer Rt ¼ 119 min, minor enantiomer Rt ¼ 120 min. 4.1.2

Procedure 2: Hydrolysis of (R)-1-Phenylethyl Octanoate O O

C7H15

OH NaOH 1M reflux overnight

4.1.2.1 • • • • • • • • • • •

O +

+ Na O

–

C7H15


(R)-1-Phenylethyl octanoate (2 g, 8 mmol) hexane (90 mL) 1 M NaOH solution (30 mL) Na2SO4, anhydrous (3 g) two-necked round-bottom flask, 25 mL capacity magnetic stirring bar hotplate with magnetic stirrer oil bath analytical balance separation funnel, 50 mL rotary evaporator.

4.1.2.3

Procedure

1. (R)-1-Phenylethyl octanoate (2 g) was heated at reflux with 1 M NaOH (30 mL) overnight. 2. The mixture was cooled to room temperature and extracted with hexane (30 mL 3). The combined organic layers were dried with anhydrous Na2SO4 and the hexane was removed by rotary evaporation. 3. The product obtained is the pure (R)-1-phenylethanol (0.70 g, 70 % yield). 1 H NMR (500 MHz, CDCl3) 7.34 7.30 (m, 5H), 4.82 (q, J ¼ 6.5, 1H), 2.34 (s, 1H), 1.45 (d, J ¼ 6.5, 3H). 13 C NMR (125 MHz, CDCl3) 145.78, 128.37 (2 C), 127.32, 125.32 (2 C), 70.21, 25.03. The purity and ee were determined by GC with a chiral column (Supelco Beta DEX 325 (90 C for 2 min, then 10 C min1 to 180 C)): major enantiomer Rt ¼ 9.11 min, minor enantiomer Rt ¼ 9.02 min.

136


Table 4.1

Effect of the size of acyl donors on the ee of the product a Acyl donor

Entry

Conversion (%)

ee (%)

O 1

CH3 O Isopropenyl acetate O

97

68

2

O CH3 Vinyl acetate O

97

65

3

C3H7 O Vinyl butanoate O

98

92

4

C7H15 O Vinyl octanoate

98

98

aReaction conditions (±)-1-Phenylethanol (0.122 g, 1 mmol), acyl donors (1.5 mmol), zeolite beta with Si/Al = 150 (50mg), Novozym 435 (30 mg) and toluene (5 mL) at 60 °C.

4.1.3

Conclusion

This DKR method can be applied to a variety of secondary alcohols. The size of the acyl donor does have a major impact on the ee of the product, as shown in Table 4.1.

References 1. Zhu, Y-.Z., Fow, K.L., Chuah, G.K. and Jaenicke, S., Dynamic kinetic resolution of secondary alcohols combining enzyme-catalyzed transesterification and zeolite-catalyzed racemisation. Chem. Eur. J. 2007, 13, 541. 2. Jaenicke, S., Chuah, G.K. and Fow, K.L., Dynamic kinetic resolution combining enzyme and zeolite catalysis. Stud. Surf. Sci. Catal. 2007, 172, 313.

4.2 Synthesis of (R)-Butyrate Esters of Secondary Alcohols by DKR

4.2

137

Synthesis of the (R)-Butyrate Esters of Secondary Alcohols by Dynamic Kinetic Resolution Employing a Bis(tetrafluorosuccinato)-bridged Ru(II) Complex S.F.G.M. van Nispen, J. van Buijtenen, J.A.J.M. Vekemans, J. Meuldijk and L.A. Hulshof

Dynamic kinetic resolution (DKR) of secondary alcohols employing Novozym 435 and a ruthenium complex as catalysts is a powerful method for the preparation of enantiomerically pure (R)-esters (Scheme 4.1).1 In this application of tandem catalysis, in situ racemization of the slow-reacting enantiomer of the alcohol enables complete conversion of the racemate into the desired enantiomer of the ester. The (R)-alcohol can be obtained by subsequent hydrolysis of the ester. Various ruthenium catalysts were successfully employed for the DKR of a broad range of secondary alcohols. Recently, we reported the DKR of a series of alcohols employing bis(tetrafluorosuccinato)-bridged Ru(II) complex (1) for the racemization (Figure 4.1, Table 4.2).2 Isopropyl butyrate was used as the acyl donor and performing the reaction at reduced pressure (200 mbar) afforded excellent yields of the (R)-butyrate esters, as was previously reported by Verzijl et al.3 employing a different ruthenium catalyst. OH R

OCOR''

lipase, acyl donor fast

R'

R

R'

Ru-catalysed racemization

OH R

OCOR''

lipase, acyl donor slow

R'

R

R'

Scheme 4.1 DKR of secondary alcohols

F

F

F

P O

O F O

Ru H

O H OC P

Ru

O

H

O F

O H

O

P CO P

O

P

= rac-BINAP

O

P F

F F

1 Figure 4.1

Bis(tetrafluorosuccinato)-bridged Ru(II) complex 1

138

DKR for the Synthesis of Esters, Amides and Acids Using Lipases Table 4.2 Dynamic kinetic resolution of various racemic alcohols. Reprinted from reference 2 with permission from Elsevier Entry

Alcohol

Time (h)

Product

Yielda,b (%)

Eea (%)

95 >99 (87)

>99 >99

O

OH

1

C3H7

O

10 24

2a 3a O OH

2d

C 3H 7

O

23

86

87

2b 3b O

OH C3H7

O

3c,e

30 F3C

96 (79)f

>99

98 (63)f

98

98

79

F3C

2c

3c O OH

4c

C3H7

O

31 O

O

2d

3d OH

O

5c

O

23

O

O

C3H7

2e 3e a

Determined by chiral gas chromatography.

b

Isolated yield in parentheses.

c

The ketone corresponding to the substrate (1.5 mmol) was added to the reaction mixture together with the substrate itself, isopropylbutyrate and toluene. d Novozym 435: 0.05 g; Ru-catalyst: 0.4 mol%. e Novozym 435: 0.4 g. f Product not separated from ketone; calculated yield.

4.2.1


General DKR procedure: OH

isopropyl butyrate (2 eq.), K2CO3, toluene, 70 °C

Ph 2

O

Novozym 435, 1 (0.1 mol%)

OH O

C3H 7

Ph 3

+

4.2 Synthesis of (R)-Butyrate Esters of Secondary Alcohols by DKR

• • • • • • • • • • • • • •

139

Novozym 435 (0.1 g) complex 1 (0.018 g, 0.0093 mmol) K2CO3 (0.5 g) substrate (9 mmol) isopropyl butyrate (18 mmol) toluene (9 mL) Schlenk tube vacuum oven P2O5 oil bath magnetic stirrer plate filter paper rotary evaporator Kugelrohr apparatus.

4.2.2

Procedure

1. Novozym 435 (0.10 g), complex 1 (0.018 g, 0.0093 mmol) and K2CO3 (0.5 g, 3.8 mmol) were dried overnight in a Schlenk tube under vacuum at 50 C in the presence of P2O5. 2. The substrate (9 mmol), isopropyl butyrate (18 mmol) and toluene (9 mL) were then added and the Schlenk tube was inserted in an oil bath at 73 C, which indicated the started of the reaction. The reaction mixture was stirred at 70 C for 23 h at a pressure of 200 mbar. Small aliquots of reaction mixture were taken for gas chromatography analysis. For preparative purposes, the reaction mixture was concentrated, filtered, washed with toluene and concentrated in vacuo to yield the crude product. 3. Further purification of the crude product by distillation in a Kugelrohr apparatus provided the (R)-esters. (R)-1-Phenylethyl butyrate (3a). Yield: 87 %. 1H NMR (300 MHz, CDCl3) (ppm) 0.95 (t, 3H, CH3), 1.55 (d, 3H, CH3), 1.63 (sextet, 2H, CH2), 2.32 (t, 2H, CH2), 5.92 (q, 1H, CH), 7.32 (5H, Ar-H). (R)-1-phenylethyl butyrate: 8.9 min. ½25 D ¼ þ91:3 (c ¼ 0.98, CHCl3). (R)--Methyl-4-(trifluoromethyl)benzyl butyrate (3c). 1H NMR (300 MHz, CDCl3) (ppm) 0.95 (t, 3H, CH3), 1.55 (d, 3H, CH3), 1.69 (sextet, 2H, CH2), 2.35 (t, 2H, CH2), 5.94 (q, 1H, CH), 7.48 (d, 2H, Ar-H-2,6), 7.61 (d, 2H, Ar-H-3,5). (R)--Methyl-4-methoxybenzyl butyrate (3d). 1H NMR (300 MHz, CDCl3) (ppm) 0.95 (t, 3H, CH3), 1.52 (d, 3H, CH3), 1.69 (sextet, 2H, CH2), 2.31 (t, 2H, CH2), 3.80 (s, 3H, CH3), 5.88 (q, 1H, CH), 6.89 (d, 2H, Ar-H-3,5), 7.32 (d, 2H, Ar-H-2,6).

References 1. Other groups’ works: (a) Larsson, A.L.E., Persson, B.A. and Ba¨ckvall, J-E., Enzymatic resolution of alcohols coupled with ruthenium-catalyzed racemization of the substrate alcohol. Angew. Chem. Int. Ed. Engl., 1997, 36, 1211. (b) Persson, B.A., Larsson, A.L.E., Le Ray, M. and Ba¨ckvall, J.-E., Ruthenium- and enzyme-catalyzed dynamic kinetic resolution of secondary alcohols. J. Am. Chem. Soc., 1999, 121, 1645. (c) Choi, J.H., Kim, Y.H., Nam, S.H., Shin, S.T., Kim, M.-J. and Park, J., Aminocyclopentadienyl ruthenium chloride: catalytic

140


racemization and dynamic kinetic resolution of alcohols at ambient temperature. Angew. Chem. Int. Ed., 2002, 41, 2373. (d) Choi, J.H., Choi, Y.K., Kim, Y.H., Park, E.S., Kim, E.J., Kim, M.-J. and Park, J., Aminocyclopentadienyl ruthenium complexes as racemization catalysts for dynamic kinetic resolution of secondary alcohols at ambient temperature. J. Org. Chem., 2004, 69, 1972. (e) Martı`n-Matute, B., Edin, M., Boga´r, K. and Ba¨ckvall, J.-E., Highly compatible metal and enzyme catalysts for efficient dynamic kinetic resolution of alcohols at ambient temperature. Angew. Chem. Int. Ed., 2004, 43, 6535. (f) Martı`n-Matute, B., Edin, M., Boga´r, K., Kaynak, F.B. and Ba¨ckvall, J-E., Combined ruthenium(II) and lipase catalysis for efficient dynamic kinetic resolution of secondary alcohols. Insight into the racemization mechanism. J. Am. Chem. Soc., 2005, 127, 8817. (g) Kim, N., Ko, S.B., Kwon, M.S., Kim, M.J. and Park, J., Air-stable racemization catalyst for dynamic kinetic resolution of secondary alcohols at room temperature. Org. Lett., 2005, 7, 4523. 2. Van Nispen, S.F.G.M., van Buijtenen, J., Vekemans, J.A.J.M., Meuldijk, J. and Hulshof, L.A., Efficient dynamic kinetic resolution of secondary alcohols with a novel tetrafluorosuccinato ruthenium complex. Tetrahedron: Asymm., 2006, 17, 2299. 3. Verzijl, G.K.M., de Vries, J.G. and Broxterman, Q.B., Removal of the acyl donor residue allows the use of simple alkyl esters as acyl donors for the dynamic kinetic resolution of secondary alcohols. Tetrahedron: Asymm., 2005, 16, 1603.

4.3 DKR of 6,7-Dimethoxy-1-methyl-1,2,3,4-tetrahydroisoquinoline

4.3

141

Dynamic Kinetic Resolution of 6,7-Dimethoxy-1-methyl-1,2,3,4tetrahydroisoquinoline Michael Page, John Blacker and Matthew Stirling

Dynamic kinetic resolution is a technique that combines a racemization with a simultaneous resolution to overcome the inherent 50 % yield limit of kinetic resolution allowing a theoretical 100 % yield. Recently, a novel chemoenzymatic system has been developed for the dynamic kinetic resolution of 6,7-dimethoxy-1-methyl-1,2,3,4-tetrahydroisoquinoline,1 building on kinetic resolution methodology developed by Breen.2 The corresponding (R)-carbamate was isolated in high yield and enantiomeric excess (Figure 4.2). MeO NH

MeO

0.2 mol% [IrCp*I2]2, 50 %w/w Candida rugosa,

+ MeO

Toluene, 40 °C, 23 hrs

MeO

Figure 4.2 Dynamic tetrahydroisoquinoline

82% yield, 96% ee

kinetic

resolution

I I

• • • • • • •

of

6,7-dimethoxy-1-methyl-1,2,3,4-

Procedure 1: Synthesis of the Amine Racemization Catalyst Pentamethylcyclopentadienyliridium(III) Iodide Dimer

Ir

4.3.1.1

O O

O

O O

4.3.1

N

MeO

Ir

I

I


Pentamethylcyclopentadienyliridium(III) chloride dimer (4.57 g) sodium iodide (8.55 g) argon cylinder acetone (95 % purity and 98.5 % enantiomeric excess.6 The conversion of acetophenone to acetophenone cyanohydrin and enantiomeric excess were determined by gas chromatographic analysis after product derivatisation as the trifluoroacetate. GC was performed using a Chiraldex capillary GC column (G-PN – g-Cyclodextrin, Propionyl) from Astec using a CP3800 (Varian) with a flame ionization detector. Carrier gas was helium at 2 mL min1. Temperature gradient: 80 °C for 0.5 min, raise at 10.8°C min1 to 130 °C and hold 130 °C for 15 min. The injector and detector temperatures were set to 250 °C.

264

Synthesis of Cyanohydrins Using Hydroxynitrile Lyases

A sample of the suspension (100 ml) was extracted with 100 ml of diisoropyl ether. 50 ml from the resulting organic phase (or if available: 1 ml crude product) were added to a mixture of 500 ml dichloromethane, 50 ml trifluoroacetic anhydride and 50 ml pyridine for the acetylation procedure. The mixture was then directly injected into the gas chromatograph. All waste solutions from the reaction were collected and disposed with hydrogen peroxide. 8.3.2

Conclusion

The use of organic-solvent-free systems can be applied to the cyanohydrin synthesis of a wide range of acetophenone derivatives (Table 8.2); electronegative substituents (e.g. fluorine) facilitate high conversions and enantiomeric excess of the product, whereas electropositive substituents (e.g. methoxy-) result in low to no conversion into the corresponding cyanohydrins.

Table 8.2

Acetophenone (AP) derivative cyanohydrin formationa

AP DERIVATIVE 0

4 -F-AP 30 -F-AP 20 -F-AP 20 ,30 ,40 ,50 ,60 -F-AP 40 -CL-AP 30 -CL-AP 20 -CL-AP 40 -BR-AP 30 -BR-AP 20 -BR-AP 40 -I-AP 20 -I-AP 40 -ME-AP 30 -ME-AP 20 -ME-AP 40 -MEO-AP 30 -MEO-AP 20 -MEO-AP 40 -NO2-AP 30 -NO2-AP 20 -NO2-AP 40 -NH2-AP 30 -NH2-AP 20 -NH2-AP 40 -OH-AP 30 -OH-AP 20 -OH-AP

TIME (H)

CONVERSIONb (%)

EE (S) (%)

1.5 3 3 6 6 6 4.5

14 48 71 99% ee >98% yield glucose

GDH-103

Figure 9.1 Production of the (S)-bis-(trifluoromethyl)phenylethanol using alcohol dehydrogenase enzyme from R. erythropolis

• • • • •

50 mM potassium phosphate buffer pH 7.2 (10 L) 3,5-bistrifluoromethylphenyl ketone (1 kg) reaction vessel with temperature and pH control 2 M sodium hydroxide heptane (10 L)

9.1.1.2

Procedure þ

1. NAD (40 g), glucose (60 g), alcohol dehydrogenase ADH-RE (3.4 g) and glucose dehydrogenase GDH-103 (3.1 g) were added to 50 mM potassium phosphate buffer pH 7.2 (10 L) that was stirring at 45 °C. The reaction was started with the addition of ketone substrate (1 kg) and aged for 24 h while maintaining a pH of 6.5 through the addition of 2 M sodium hydroxide. 2. The reaction was extracted twice with 5 L heptane. The organic layers were then combined, washed with 2.5 L water and evaporated by distillation until the alcohol product concentration was 200 g L1. The solution was cooled to 35 °C and seeded with 1 g alcohol product prior to aging for 1 h and then cooling again to 10 °C. The alcohol crystallized into a solid with >99% purity. 1

H NMR: 7.85 (s, 2H), 7.80 (s, 1H), 5.05 (qd, J ¼ 6.5, 3.3, 1H), 2.04 (d, J ¼ 3.3, 1H), 1.56 (d, J ¼ 6.5, 3H). 13C NMR: 148.44, 131.99 (q, J ¼ 33.2), 125.87 (br q, J ¼ 2.8), 123.58 (q, J ¼ 272.6), 121.53 (septet, J ¼ 3.9), 69.31, 25.79. Chiral analysis for ee determination was by normal-phase high-performance liquid chromatography with a Chiralcel OD-H column using 98 % hexanes/2 % 2-propanol at 1 mL min1, 25 °C and monitoring at 265 nm. 9.1.2

Conclusion

This novel biocatalytic method for the production of (S)-bis(trifluoromethyl)phenylethanol was easily and reproducibly demonstrated up to pilot plant scale in reactions generating 25 kg of >99% ee material. Substrate concentrations were increased up to 580 mM,

9.1 Asymmetric Synthesis of (S)-Bis(trifluoromethyl)phenylethanol

275

resulting in a space–time yield of 260 g L1 day1. Additionally, enantiocomplementary results were obtained by using an identical procedure with the commercially available isolated ketoreductase KRED-101 (Biocatalytics) in place of alcohol dehydrogenase ADH-RE.

Reference 1. Pollard, D., Truppo, M., Pollard, J., Chen, C. and Moore, J. , Effective synthesis of (S)-3,5bistrifluoromethylphenyl ethanol by asymmetric enzymatic reduction. Tetrahedron Asymm., 2006, 17, 554–559.

276

Synthesis of Chiral sec-Alcohols by Ketone Reduction

9.2

Enantioselective and Diastereoselective Enzyme-catalyzed Dynamic Kinetic Resolution of an Unsaturated Ketone Birgit Kosjek, David Tellers and Jeffrey Moore

Whole-cell cultures and isolated enzymes have been shown to be very useful in catalyzing highly chemoselective reductions of ,-unsaturated ketones. The presence of an additional racemic centre in a ketone substrate for this reduction has a strong potential for decreasing the overall yield of the reaction by introducing a competing directing effect on the enzyme. To reduce such a compound effectively and efficiently, a biocatalytic process was developed that incorporates a racemization step to increase the theoretical yield of enantiomerically pure product to 100 %.1 This process was used to generate allylic alcohol with an enantioselectivity of 95 % ee and a diastereoselectivity of 99 % de (Figure 9.2). O

OH KRED-108 CO2Me

CO2Me

CO2Me 1

99% de 95% ee 94% yield

CO2Me NADPH

NADP+

2

OH

O KRED-104

Figure 9.2

9.2.1 9.2.1.1 • • • • • • •

Enantioselective and diastereoselective reduction of a,-unsaturated ketones

Procedure 1: Ketoreductase Reduction of Ketone 1 Materials and Equipment

Ketoreductase KRED-104 (Codexis Inc, 120 mg) ketoreductase KRED-108 (Codexis Inc, 30 mg) nicotinamide adenine dinucleotide phosphate (NADPþ, 30 mg) 0.5 M potassium phosphate buffer pH 6.5 (9.5 mL) ketone substrate (100 mg) isopropanol (0.5 mL) ethyl acetate (10 mL).

9.2.1.2

Procedure

1. Ketoreductase enzymes KRED-104 (120 mg) and KRED-108 (30 mg) and NADPþ (30 mg) were added to 0.5 M potassium phosphate buffer pH 6.5 (9.5 mL) that was stirring at 35 °C. The reaction was started with the addition of a solution of ketone substrate (100 mg) in isopropanol (0.5 mL) and aged for 12 h.

9.2 Enzyme-catalyzed Dynamic Kinetic Resolution of an Unsaturated Ketone

277

2. The reaction was extracted with an equal volume of ethyl acetate and the organic layer was evaporated, resulting in isolation of the product 2 at a yield of 94 %, a de of 99% cis, and an ee of 95 % in favour of the S-enantiomer. 1

H NMR (399.9 MHz; acetonitrile-d3, 27 °C) 6.95 (s, 1H), 4.30 (m, 1H), 3.65 (s, 3H), 3.42 (m, 1H), 2.45 (m, 2H), 2.35 (m, 2H), 2.1 (m, 2H) 1.98 (m, 1H). 13C NMR (125 MHz, tetrahydrofuran-d8, 27 °C) ¼ 24.8, 28.9, 41.0, 52.7, 52.8, 66.3, 130.0, 144.4, 167.6, 175.1 ppm. Conversion and diastereomeric excess were determined on an Agilent HPLC system using a Zorbax eclipse XDB C18 column (4.6 mm 150 mm) at a gradient from 35/65 MeCN/water (0.1 % H3PO4) to 95/5 over 14 min at 1 mL min1, room temperature, 210 nm. Enantiomeric excess was determined with a Berger SFC system employing a tandem Chiralpak OD (250 mm 4.6 mm)–Chiralpak OB (250 mm 4.6 mm), isocratic 3 % 2-propanol/CO2 at 2 mL min1, 200 bar, 35 °C, 30 min. Alternatively, product enantiomeric excess could be measured by chiral gas chromatography: Agilent GC system, Varian Chiralsil-Dex Cb (25 m 0.32 mm, 0.25 mm film thickness) ramp from 70 °C to 190 °C at 2 °C min1, ramp to 200 °C at 1 °C min1, hold for 10 min, average velocity 39 cm s1. 9.2.2

Conclusion

Whereas the use of chemical catalysts to reduce this unsaturated ketone does not afford any diastereoselective discrimination, the biocatalytic method described here generates product that is almost exclusively the cis diastereomer at a very high ee of 95 %. Important process improvements included optimizing the ester moiety of the starting material, after it was found to have a significant impact on the observed enantioselectivity, and the inclusion of isopropanol in the reaction mixture to serve both as the hydrogen source for the recycling of the cofactor NADPH and as a cosolvent for increasing the solubility of the ketone substrate.

Reference 1. Kosjek, B., Tellers, D.M., Biba, M., Farr, R. and Moore, J.C., Biocatalytic and chemocatalytic approaches to the highly stereoselective 1,2-reduction of an ,-unsaturated ketone. Tetrahedron Asymm., 2006, 17, 2798–2803.

278


9.3

Enzyme-catalysed Synthesis of a-Alkyl-b-hydroxy Ketones and Esters by Isolated Ketoreductases Ioulia Smonou and Dimitris Kalaitzakis

Using isolated enzymes as catalysts for organic reactions is becoming a more standardized and practical tool in the hands of organic chemists.1 The biocatalytic reduction of -alkyl-1,3diketones and -alkyl--keto esters employing commercially available reduced nicotinamide adenine dinucleotide phosphate (NADPH)-dependent ketoreductases (KREDs) proved to be a highly efficient method for the preparation of optically pure keto alcohols, 1,3-diols or hydroxy esters.2,3 These enzymatic reactions provide a simple, highly stereoselective and quantitative method for the synthesis of different stereoisomers of valuable chiral synthons from nonchiral, easily accessible 1,3-diketones or keto esters (Figure 9.3). Chiral keto alcohols and diols represent very useful synthons in organic synthesis and have been used as precursors in the synthesis of various biologically active compounds4,5 and pharmaceuticals. OH KRED-102

3

4

O 2

1

3R,4S > 99%ee, > 99%de

1

3S,4R > 99%ee, > 98%de

NADPH O

O

OH KRED-A1B 4

NADPH

O

3 2

OH KRED-108

4

O 3

2

NADPH

1

3S,4S > 99%ee, > 98%de

Figure 9.3 Enzyme-catalysed stereoselective reduction of 3-allyl-2,4-pentanedione

9.3.1

Procedure 1: Synthesis of (3R,4S)-3-Allyl-4-hydroxy-2-pentanone OH

9.3.1.1 • • • • • • • •

O


200 mM phosphate buffer solution, pH 6.9 (100 mL) 3-allyl-2,4-pentanedione (700 mg, 50 mmol) glucose (2.16 g, 120 mmol) NADPH (45 mg) glucose dehydrogenase (50 mg) KRED-102 (50 mg) (Codexis Inc.) NaOH solution (2 M) ethyl acetate (200 mL)

9.3 Synthesis of a-Alkyl--hydroxy Ketones and Esters by Isolated KREDS

• • • • • • •

279

saturated NaCl solution (70 mL) anhydrous MgSO4 (3 g) pH meter one-necked reaction flask equipped with magnetic stirring bar, 250 mL magnetic stirring plate one 250 mL separatory funnel rotary evaporator.

9.3.1.2

Procedure

1. A 200 mM phosphate-buffered solution, pH 6.9 (100 mL), containing 3-allyl2, 4-pentanedione (700 mg, 50 mmol), glucose (2.16 g, 120 mmol), NADPH (45 mg), glucose dehydrogenase (50 mg) and KRED-102 (50 mg) was stirred at room temperature for 24 h, until gas chromatography (GC) analysis of the crude extracts showed complete reaction. Periodically, the pH was readjusted to 6.9 with NaOH (2 M). 2. The product (3R,4S)-3-allyl-4-hydroxy-2-pentanone was isolated by extracting the crude reaction mixture with EtOAc (2 100 mL). The combined organic layers were then extracted with saturated NaCl solution (70 mL), dried over MgSO4 and evaporated to dryness to afford optically active (3R,4S)-3-allyl-4-hydroxy-2-pentanone (617 mg, 87 %). 1

H NMR (CDCl3; 500 MHz) 5.74–5.82 (m, 1H), 5.01–5.11 (m, 2H), 4.01–4.07 (m, 1H), 2.64–2.68 (m, 1H), 2.39–2.42 (m, 2H), 2.18 (s, 3H), 1.18 (d, J ¼ 6.5 Hz, 3H). The optical purity was determined by chiral GC, using a 20 % permethylated cyclodextrin column, after esterification of the pure product with (CF3CO)2O in dry CH2Cl2 (65 °C isothermal; carrier gas: N2, pressure 70 kPa). TR ¼ 26.305 min (>99%, (3R,4S)-3-allyl-4hydroxy-2-pentanone). The enantiomeric purity was estimated to be >99 % and the diastereomeric purity >99 %. 9.3.2

Procedure 2: Synthesis of (3S,4R)-3-Allyl-4-hydroxy-2-pentanone OH

9.3.2.1 • • • • • • • •

O


200 mM phosphate buffer solution, pH 6.9 (100 mL) 3-allyl-2,4-pentanedione (700 mg, 50 mmol) glucose (2.16 g, 120 mmol) NADPH (45 mg) glucose dehydrogenase (50 mg) KRED-A1B (50 mg) (Codexis Inc.) NaOH solution (2 M) ethyl acetate (200 mL)

280

• • • • • • •


saturated NaCl solution (70 mL) anhydrous MgSO4 (3 g) pH meter one-necked reaction flask equipped with magnetic stirring bar, 250 mL magnetic stirring plate one 250 mL separatory funnel rotary evaporator.

9.3.2.2

Procedure

1. A 200 mM phosphate-buffered solution, pH 6.9 (100 mL), containing 3-allyl-2, 4-pentanedione (700 mg, 50 mmol), glucose (2.16 g, 120 mmol), NADPH (45 mg), glucose dehydrogenase (50 mg) and KRED-A1B (50 mg) was stirred at room temperature for 8 h, until GC analysis of crude extracts showed complete reaction. Periodically, the pH was readjusted to 6.9 with NaOH (2 M). 2. Product (3S,4R)-3-allyl-4-hydroxy-2-pentanone was isolated by extracting the crude reaction mixture with EtOAc (2 100 mL). The combined organic layers were then extracted with saturated NaCl solution (70 mL), dried over MgSO4 and evaporated to dryness to afford optically active (3S,4R)-3-allyl-4-hydroxy-2-pentanone (604 mg, 85 %). 1

H NMR (CDCl3; 500 MHz) 5.74–5.82 (m, 1H), 5.02–5.12 (m, 2H), 4.02–4.07 (m, 1H), 2.64–2.68 (m, 1H), 2.39–2.43 (m, 2H), 2.18 (s, 3H), 1.18 (d, J ¼ 6.5 Hz, 3H). The optical purity was determined by chiral GC, using a 20 % permethylated cyclodextrin column, after esterification of the pure product with (CF3CO)2O in dry CH2Cl2 (65 °C isothermal; carrier gas: N2, pressure 70 kPa). TR ¼ 27.604 min (99 %, (3R,4S)-3-allyl-4-hydroxy-2-pentanone), TR ¼ 28.776 min (1 %, (3R,4R)-3allyl-4-hydroxy-2-pentanone). The enantiomeric purity was estimated to be >99 % and the diastereomeric purity 98 %. 9.3.3

Procedure 3: Synthesis of (3S,4S)-3-Allyl-4-hydroxy-2-pentanone OH

9.3.3.1 • • • • • •

O


200 mM phosphate buffer solution, pH 6.9 (100 mL) 3-allyl-2,4-pentanedione (700 mg, 50 mmol) glucose (2.16 g, 120 mmol) NADPH (45 mg) glucose dehydrogenase (50 mg) KRED-108 (70 mg) (Codexis Inc.)


• • • • • • • • • •

281

NaOH solution (2 M) ethyl acetate (200 mL) saturated NaCl solution (70 mL) anhydrous MgSO4 (3 g) pH meter one-necked reaction flask equipped with magnetic stirring bar, 250 mL magnetic stirring plate filter paper one 250 mL separatory funnel rotary evaporator.

9.3.3.2

Procedure

1. A 200 mM phosphate-buffered solution, pH 6.9 (100 mL), containing 3-allyl-2, 4-pentanedione (700 mg, 50 mmol), glucose (2.16 g, 120 mmol), NADPH (45 mg), glucose dehydrogenase (50 mg) and KRED-108 (70 mg) was stirred at room temperature for 24 h, until GC analysis of crude extracts showed complete reaction. Periodically, the pH was readjusted to 6.9 with NaOH (2 M). 2. Product (3S,4S)-3-allyl-4-hydroxy-2-pentanone was isolated by extracting the crude reaction mixture with EtOAc (2 100 mL). The combined organic layers were then extracted with saturated NaCl solution (70 mL), dried over MgSO4 and evaporated to dryness to afford optically active (3S,4S)-3-allyl-4-hydroxy-2-pentanone (614 mg, 86 %). 1

H NMR (CDCl3; 500 MHz) 5.67–5.76 (m, 1H), 5.03–5.11 (m, 2H), 3.92–3.97 (m, 1H), 2.60–2.65 (m, 1H), 2.34–2.37 (m, 2H), 2.19 (s, 3H), 1.23 (d, J ¼ 6 Hz, 3H). The optical purity was determined by chiral GC, using a 20 % permethylated cyclodextrin column, after esterification of the pure product with (CF3CO)2O in dry CH2Cl2 (65 °C isothermal; carrier gas: N2, pressure 70 kPa). TR ¼ 26.739 min (1 %, (3R,4S)-3-allyl-4hydroxy-2-pentanone), TR ¼ 29.864 min (99 %, (3S,4S)-3-allyl-4-hydroxy-2-pentanone). The enantiomeric purity was estimated to be >99 % and the diastereomeric purity 98 %.

O

O

R1

R2 R 3 R4

OH

KRED R1 NADPH NADP+

I

O

* *

R2

R3 R 4 II

GDH Gluconolactone

Figure 9.4

D-Glucose

Enzymatic reduction of a-alkyl-1,3-diketones with NADPH-dependent KREDs

282


Table 9.1 Enzyme-catalysed stereoselective reduction of diketones/keto esters to keto alcohols/hydroxy esters Entry R1

R2

R3

R4

KRED

Product yield (%) A R-Alkyl, S-OH

B S-Alkyl, S-OH

C S-Alkyl, R-OH

D R-Alkyl, R-OH

>99 3R,4S 3

—

—

—

94 3S,4S —

—

3

—

—

>99 [12 h]

—

95 3S,4R —

5

>99 [1 h]

—

>99 [24 h]

97 4S,5R —

3

>99 [40 min]

—

>99 [12 h]

—

—

>99 [24 h]

96 2S,3S —

—

1

2

—

>99 [1 h]

—

10

>99 [24 h]

>99 3S,4S —

—

90 3R,4R —

>99 [6 h]

—

—

>99 [24 h]

15

—

85 2S,3S

>99 [6 h]

1

Me Me Me H

102

2

Me Me Me H

127

3

Me Me Et

H

102

4

Me Me Et

H

A1B

>99 3R,4S —

5

Me Me Et

H

118

—

6

Et

Et

Me H

A1B

—

7

Et

Et

Me H

119

99 2S,3R 3

10

Me Me Me Allyl 101

11

Me Me Me Allyl A1B

98 3R,4S —

12

Me Me Me Allyl 118

—

13

Me OEt Me H

102

14

Me OEt Me H

107

>99 2R,3S —

9.3.4

Conversion (%) [time]

>98 3S,4S >99 4S,5S —

>99 [24 h] 90 [24 h]

92 [24 h]

Conclusion

The enzymatic transformation of a large number of -monoalkyl and dialkyl symmetrical and nonsymmetrical 1,3-diketones and keto esters (Figure 9.4) shows excellent chemical and optical yield and can be tailored to afford most of the four possible diastereomers from the same starting substrate at will, depending on the chosen enzyme. Besides being regio- and stereo-selective, these enzymes exhibited high chemoselectivity by giving a keto alcohol or hydroxy ester and not the diol. Table 9.1 shows some examples of the different substrates that can be reduced to various single diastereomers of the same compound, as the keto alcohols and hydroxy esters, by choosing the use of different enzymes. The chemoenzymatic syntheses of the aggregation pheromones (þ)-Sitophilure and Sitophilate by the use of the above isolated, NADPHdependent KREDs were successfully accomplished by our group4,5 with high chemical and optical purities (98 % de, >99 % ee).


283

References 1. Faber, K., Biotransformations in Organic Chemistry. 1997, Springer-Verlag, Berlin, pp. 160– 206. 2. Kalaitzakis, D., Rozzell, J.D., Kambourakis, and S. Smonou, I., Highly stereoselective reductions of -alkyl-1,3-diketones and -alkyl--keto esters catalyzed by isolated NADPH-dependent ketoreductases. Org. Lett., 2005, 7, 4799–4801. 3. Kalaitzakis, D., Rozzell, J.D., Kambourakis, S. and Smonou, I., Synthesis of valuable chiral intermediates by isolated ketoreductases: application in the synthesis of -alkyl--hydroxy ketones and 1,3-diols. Adv. Synth. Catal., 2006, 348, 1958–1969. 4. Kalaitzakis, D., Rozzell, J.D., Kambourakis, S. and Smonou, I., A two-step chemoenzymatic synthesis of the natural pheromone (þ)-Sitophilure utilizing isolated, NADPH-dependent ketoreductases. Eur. J. Org. Chem., 2006, 2309–2313. 5. Kalaitzakis, D., Kambourakis, S., Rozzell, J.D. and Smonou, I., Stereoselective chemoenzymatic synthesis of sitophilate: a natural pheromone. Tetrahedron Asymm. 2007, 18, 2418–2426.

284


9.4

Asymmetric Reduction of Phenyl Ring-containing Ketones Using Xerogel-encapsulated W110A Secondary Alcohol Dehydrogenase from Thermoanaerobacter ethanolicus Musa M. Musa, Karla I. Ziegelmann-Fjeld, Claire Vieille, J. Gregory Zeikus and Robert S. Phillipsa

There has been a growing interest in using enzymes for asymmetric transformations of unnatural organic compounds in organic solvents.1 Recently, we have used xerogelimmobilized W110A mutant secondary alcohol dehydrogenase from Thermoanaerobacter ethanolicus (W110A TeSADH) to reduce a series of phenyl ringcontaining ketones to the corresponding (S)-alcohols in good yields and high optical purities in organic solvents (Figure 9.5).2 The resulting alcohols have (S)-configuration, in agreement with Prelog’s rules, in which the reduced nicotinamide adenine dinucleotide phosphate (NADPH) cofactor transfers its pro-R hydride to the re face of the ketone. 9.4.1

Procedure 1: Preparation of Xerogel-encapsulated W110A TeSADH

9.4.1.1 • • • • • • • •


Tetramethyl orthosilicate (TMOS, 2.10 g) distilled water (0.47 g) HCl (0.04 m, three drops) W110A TeSADH (0.43 mg) NADPþ (3.0 mg, 3.6 mmol) tris-HCl buffer (50 mM, pH 7.0, 1.0 mL) one 10 mL round-bottomed flask sonifier.

9.4.1.2

Procedure

1. The silica sol was prepared by mixing TMOS (2.10 g), distilled water (0.47 g) and HCl (0.04 M, three drops). The mixture was then sonicated until one layer was formed.

O

Hexane

O

OH

xerogel W110A TeSADH R na

NADPH

NADP+

R (S )-nb

6a OH

O

OH xerogel W110A TeSADH

(S)-6b

R = Ph(CH2)2, PhOCH2, p-MeOC6H4(CH2)2, PhCH2, p-MeOC6H4CH2

Figure 9.5 Reduction of ketones with W110A secondary alcohol dehydrogenase

9.4 Asymmetric Reduction of Phenyl Ring-containing Ketones

285

2. The gels were prepared by mixing the above sol (1.0 mL) with enzyme stock (1.0 mL) in a 10 mL round-bottomed flask. The enzyme stock was prepared in 50 mM tris-HCl buffer (pH 8.0) such that the concentration of the enzyme, expressed and purified as described previously,3 was 0.43 mg mL1, and that of NADPþ was 3.0 mg mL1. The sol–gel was then left in the same flask sealed with Parafilm at room temperature for 48 h to allow gel to age. 3. The hydrogel was dried at room temperature in air for 24 h to give hydrated silica, SiO2nH2O, the so-called xerogel. 9.4.2 9.4.2.1 • • • • • • • • • • • • •

Procedure 2: Asymmetric Reduction Using Xerogel-encapsulated W110A TeSADH in Organic Solvents Materials and Equipment

Ketone substrate (0.34 mmol) 2-propanol (600 mL) hexane (2 mL) ethyl acetate (4 mL) anhydrous Na2SO4 pyridine acetic anhydride ˚ , 32–63 mm) silica gel (60 A one 10 mL round-bottomed flask equipped with a magnetic stirrer hot and magnetic stirrer plate filter paper rotary evaporator equipment for column chromatography.

9.4.2.2

Procedure

1. All reactions were performed using W110A TeSADH (0.43 mg) and NADPþ (3.0 mg, 3.6 mmol) encapsulated in sol–gel, substrate (0.34 mmol), 2-propanol (600 mL), and 2.0 mL of hexane in a 10 mL round-bottomed flask equipped with a magnetic stirrer. The reaction mixture was stirred at 50 °C for 12 h. 2. The sol–gel was then removed by filtration and washed with ethyl acetate (2 2 mL). The combined organic filtrates were dried with Na2SO4 and then concentrated under vacuum. 3. The remaining residue was analyzed by gas chromatography (GC) to determine the yield, then purified by silica-gel column chromatography (eluent: ethyl acetate: hexane, 15:85). The product alcohol was then converted to the corresponding acetate derivative.4 The ee was determined by GC equipped with a flame-ionization detector and a Supelco -Dex 120 chiral column (30 m, 0.25 mm (internal diameter), 0.25 mm film thickness) by using He as the carrier gas. The injector temperature was 250 °C and the detector temperature was 300 °C. The flow rate was 19.0 psi. The column was programmed between 120 °C and 170 °C.

286


Table 9.2 Asymmetric reduction of phenyl ring-containing ketones by TeSADH using Procedure 2 Entry

R

1a 2b 3c 4d 5e 6f

Ph(CH2)2 PhOCH2 p-MeOC6H4(CH2)2 PhCH2 p-MeOC6H4CH2 2-Tetralol (see Figure 9.1)

Product Yield (%)

Ee (%)

74 >99 61 80 67 94

97 >99 94 69 >99 76

20 a (S)-4-Phenyl-2-butanol: []D ¼ þ16.5 (c ¼ 1.81, CHCl3), >99 % ee, lit.5 []D 20¼ þ17.4 c ¼ 1.80, CHCl3), 99 % ee. Spectral data were consistent with that reported previously.6 20 7 b (S)-Phenoxy-2-propanol: []D ¼ þ30.7 (c ¼ 1.32, CHCl3), >99 % ee, lit. []20 D ¼ þ28.9 c ¼ 1.10, CHCl3), 99 % ee. Spectral data were consistent with that reported previously.8 20 20 c (S)-4-(4-Methoxyphenyl)-2-butanol: []D ¼ þ12.8 (c ¼ 2.41, CHCl3), >91 % ee, lit.9 []D ¼ þ30.9 c ¼ 1.0, CHCl3), 94 % ee. Spectral data were consistent with that reported previously.9 20 25 10 d (S)-1-Phenyl-2-propanol: []D ¼ þ14.5 (c ¼ 1.04, CHCl3), >37 % ee, lit. []D ¼ þ42.2 c ¼ 1.0, CHCl3), >99 % ee. Spectral data were consistent with that reported previously.11 e 20 20 (S)-4-(4-Methoxyphenyl)-2-propanol: []D ¼ þ16.3 (c ¼ 1.86, CHCl3), >99 % ee, lit.12 []D ¼ þ27.0 c ¼ 4.40, CHCl3), 95 % ee. Spectral data were consistent with that reported previously.10 f 20 20 13 (S)-2-Tetralol: []D ¼ 43.77 (c ¼ 0.911, CHCl3), >71 % ee, lit. []D ¼ 29.6 c ¼ 0.50, CHCl3), 85 % ee. Spectral data were consistent with that reported previously.14

9.4.3

Conclusion

This method allows the asymmetric reduction of hydrophobic ketones in high yields and enantioselectivities (Table 9.2). It is a facile method, not only for making the enzyme accessible to a wide variety of water-insoluble substrates by switching the traditional aqueous medium to organic media, but also for reusing the enzyme. This method allows for the use of high concentrations of substrate and catalytic quantities of cofactor, both of which are crucial for large-scale synthetic applications. Reusable catalysts for chemo-, regio-, and enantio-selective asymmetric reduction may be of industrial interest.

References 1. Faber, K., Biotransformations in Organic Chemistry, 5th edn. Springer: Heidelberg, 2004. 2. Musa, M., Ziegelman-Fjeld, K., Vieille, C., Zeikus, J. and Phillips, R., Xerogel-encapsulated W110A secondary alcohol dehydrogenase from Thermoanaerobacter ethanolicus performs asymmetric reduction of hydrophobic ketones in organic solvents. Angew. Chem. Int. Ed., 2007, 46, 3091–3094. 3. Ziegelman-Fjeld, K., Musa, M., Phillips, R., Zeikus, J. and Vieille, C., A Thermoanaerobacter ethanolicus secondary alcohol dehydrogenase mutant derivative highly active and stereoselective on phenylacetone and benzylacetone. Protein Eng. Des. Sel., 2007, 20, 47–55. 4. Ghanem, A. and Schuring, V., Lipase-catalyzed access to enantiomerically pure (R)- and (S)-trans-4-phenyl-3-butene-2-ol. Tetrahedron Asymm., 2003, 14, 57–62. 5. Nakamura, K., Inoue, Y., Matsuda, T. and Misawa, I., Stereoselective oxidation and reduction by immobilized Geotrichum candidum in an organic solvent. J. Chem. Soc. Perkin Trans. 1, 1999, 2397–2402.

9.4 Asymmetric Reduction of Phenyl Ring-containing Ketones

287

6. Kuwano, R., Uemura, T., Saitoh, M. and Ito, Y., A trans-chelating bisphosphine possessing only planar chirality and its application to catalytic asymmetric reactions. Tetrahedron Asymm.,2004, 15, 2263–2271. 7. Nakamura, K., Takenaka, K., Fujii, M. and Ida, Y., Asymmetric synthesis of both enantiomers of secondary alcohols by reduction with a single microbe. Tetrahedron Lett., 2002, 43, 3629–3631. 8. Dragovich, P. S., Prins, T. J. and Zhou, R., Palladium catalyzed, regioselective reduction of 1,2-epoxides by ammonium formate. J. Org. Chem., 1995, 60, 4922–4924. 9. Donzelli, F., Fuganti, C., Mendozza, M., Pedrocchi-Fantoni, G., Servi, S. and Zucchi, G., On the stereochemistry of the Baeyer–Villiger degradation of arylalkylketones structurally related to raspberry ketone by Beauveria bassiana. Tetrahedron Asymm., 1996, 7, 3129–3134. 10. Erde´lyi, B., Szabo´, A., Seres, G., Birincsik, L., Ivanics, J., Szatzker, G. and Poppe, L., Stereoselective production of (S)-1-aralkyl- and 1-arylethanols by freshly harvested and lyophilized yeast cells. Tetrahedron Asymm., 2006, 17, 268–274. 11. Ley, S. V., Mitchell, C., Pears, D., Ramarao, C., Yu, J. and Zhou, W., Recyclable polyureamicroencapsulated Pd(0) nanoparticles: an efficient catalyst for hydrogenolysis of epoxides. Org. Lett., 2003, 5, 4665–4668. 12. Ferraboschi, P., Grisenti, P., Manzocchi, A. and Santaniello, E., Baker’s yeast-mediated preparation of optically active aryl alcohols and diols for the synthesis of chiral hydroxy acids. J. Chem. Soc. Perkin Trans. 1, 1990, 2469–2474. 13. Stampfer, W., Kosjek, B., Faber, K. and Kroutil, W., Biocatalytic asymmetric hydrogen transfer employing Rhodococcus ruber DSM 44541. J. Org. Chem., 2003, 68, 402–406. 14. Orsini, F., Sello, G., Travaini, E. and Di Gennaro, P., A chemoenzymatic synthesis of (2R)-8substituted-2-aminotetralins. Tetrahedron Asymm., 2002, 13, 253–259.

288


9.5

(R)- and (S)-Enantioselective Diaryl Methanol Synthesis Using Enzymatic Reduction of Diaryl Ketones Matthew Truppo, Krista Morley, David Pollard and Paul Devine

The asymmetric formation of industrially useful diaryl methanols can be realized through either the addition of aryl nucleophiles to aromatic aldehydes or the reduction of diaryl ketones.1 The latter route is frequently the more desirable, as the starting materials are often inexpensive and readily available and nonselective background reactions are not as common. For good enantioselectivity, chemical catalysts of diaryl ketone reductions require large steric or electronic differentiation between the two aryl components of the substrate and, as a result, have substantially limited applicability.2,3 In contrast, recent work has shown commercially available ketoreductase enzymes to have excellent results with a much broader range of substrates in reactions that are very easy to operate (Figure 9.6).4 9.5.1

Procedure 1: General Procedure for the Ketoreductase Reduction of Diaryl Ketones

9.5.1.1 • • • • • • • • • • • •

Materials and equipment

Ketone substrate (1 g) glucose (800 mg) nicotinamide adenine dinucleotide phosphate (NADPþ, 40 mg) 0.1 M potassium phosphate buffer pH 7 (36 mL) tetrahydrofuran (THF, 4 mL) ketoreductase enzyme (Codexis Inc, 80 mg) glucose dehydrogenase enzyme (Codexis Inc, 80 mg) 2-butanone (80 mL) nitrogen gas flask (100 mL) separatory funnel rotary evaporator.

9.5.1.2

Procedure

1. Glucose (800 mg), NADPþ (40 mg), ketone substrate example (1 g) and THF (4 mL) were added to 0.1 M potassium phosphate buffer pH 7 (36 mL) that was stirring at 30 °C.

O Ar1

ketoreductase Ar2

1

Ar NADPH

O

OH 2

* Ar

Ar1

ketoreductase Ar2

Ar NADPH

NADP+ glucose

glucose dehydrogenase

Figure 9.6

* Ar2

NADP+ OH

O gluconolactone

OH 1

ketoreductase

Asymmetric reduction of diaryl ketones with ketoreductases

9.5 (R)- and (S)-Enantioselective Diaryl Methanol Synthesis

289

The reaction was started with the addition of ketoreductase (80 mg) and glucose dehydrogenase (80 mg) enzymes. 2. The reaction was extracted with 2-butanone (80 mL) that was then washed twice with 5 mL water. The organic layer was evaporated under nitrogen, yielding the alcohol product.

Table 9.3

Reduction of various diaryl ketones with ketoreductases. (R)-Alcohol

Ketone

Ee (%) Ketoreductasea R1 ¼ o-CH3 R1 ¼ m-CH3 R1 ¼ p-CH3 R1 ¼ m-NO2 R1 ¼ p-NO2 R1 ¼ o-OH R1 ¼ m-OH R1 ¼ p-OH R1 ¼ o-NH2 R1 ¼ p-OMe R1 ¼ p-NO2 R1 ¼ o-Cl R1 ¼ m-Cl R1 ¼ p-Cl R1 ¼ m-CN R2 ¼ p-Cl R1 ¼ m-CO2Me R2 ¼ p-Cl

119 CDX P2C12 119 108 119

13 55 64 51 64 99 99

119 117 114 119 119 118 108

90

108

115

99

108

97

101

77

119

O

82

101

38

120

O

44

Lactobacillus kefir

99

119

94

124

60

119

R2

R1

O

121 CDX P1H10 CDX P1H10 111 CDX P1H10 111 CDX P1H10 CDX P1H10 101 111 101 121 CDX P1H10 CDX P1H10 112

33

Ee (%) Ketoreductasea 95 92 9 99 97

O

98 99 99 34 99 84 82 96 91 60 70 64 97 99 84

(S)-Alcohol

N

N

N

O N Cl a

Ketoreductases identified by numbers refer to enzymes commercially available from Codexis (Pasadena, CA – formerly Biocatalytics) and those with CDX prefixes refer to enzymes obtained under license from Codexis (Redwood City, CA – CodexTM KRED Panel v 1.0).

290


9.5.2 9.5.2.1 • • • • • • • • • •

Procedure 2: Screening of Ketoreductase Enzymes Materials and equipment

Ketone THF NADPþ 0.1 M potassium phosphate buffer, pH 7 aqueous glucose solution ketoreductase enzyme (Codexis Inc) glucose dehydrogenase enzyme (Codexis Inc) methyl tert-butyl ether (MTBE) 96-well plate chiral analytical method.

9.5.2.2

Procedure

1. To rapidly screen libraries of ketoreductase enzymes in parallel against ketone starting materials of interest, a substrate solution containing 20 mg mL1 ketone in THF, a cofactor solution containing 5 mg mL1 NADPþ in 0.1 M potassium phosphate buffer pH 7, and a glucose solution containing 20 mg mL1 glucose in water were prepared. 2. For each Biocatalytics (Codexis Pasadena) ketoreductase enzyme, 50 mL each of the substrate, cofactor and glucose solutions were added to 350 mL 0.1 M potassium phosphate buffer pH 7, 1 mg ketoreductase and 1 Mg glucose dehydrogenase enzymes in one location of a 96-well plate. 3. For each Codexis ketoreductase enzyme, 50 mL each of the substrate and cofactor solutions were added to 300 mL isopropanol, 100 mL 0.1 M potassium phosphate buffer pH 7 and 1 mg ketoreductase in one location of a 96-well plate. 4. After aging the reactions for 24 h at 30 °C, they were each extracted with 1 mL MTBE for analysis using a suitable chiral method. 9.5.3

Conclusion

A large number of diaryl ketone substrates, including those listed in Table 9.3, have been reduced with high enantioselectivity with the protocol described here. Unlike analogous chemical catalysts, the commercially available biocatalysts displayed no dependence on ortho substitutions or electronic dissymmetry, and produced diaryl methanols with good to excellent ee values in nearly all cases.

References 1. Devaux-Basseguy, R., Bergel, A. and Comtat, M., Potential applications of NAD(P)-dependent oxidoreductases in synthesis: a survey. Enzyme Microb. Technol., 1997, 20, 248. 2. Welch, C. J., Grau, B., Moore, J. and Mathre, D., Use of chiral HPLC–MS for rapid evaluation of the yeast-mediated enantioselective bioreduction of a diaryl ketone. J. Org. Chem., 2001, 66, 6836. 3. Itsuno, S. Enantioselective reduction of ketones. In Organic Reactions, vol. 52, Paquette, L.A. (ed.), John Wiley & Sons, Inc.: New York, 1998, pp. 395–576. 4. Truppo, M. D., Pollard, D. and Devine, P., Enzyme-catalyzed enantioselective diaryl ketone reductions. Org. Lett., 2007, 9, 335.

9.6 Enantioselective and Efficient Synthesis of Methyl (R)-o-Chloromandelate

9.6

291

Highly Enantioselective and Efficient Synthesis of Methyl (R)-oChloromandelate, Key Intermediate for Clopidogrel Synthesis, with Recombinant Escherichia coli Tadashi Ema, Nobuyasu Okita, Sayaka Ide and Takashi Sakai

Clopidogrel is a platelet aggregation inhibitor widely administered to atherosclerotic patients with the risk of a heart attack or stroke that is caused by the formation of a clot in the blood. Worldwide sales of Plavix (clopidogrel bisulfate) amounted to $6.4 billion per year (data for the 12 months ending June 2006), which ranks second to Lipitor for sales.1 We have recently found that methyl (R)-o-chloromandelate ((R)-1), which is a key intermediate for clopidogrel synthesis, can be obtained in >99 % ee by the asymmetric reduction of methyl o-chlorobenzoylformate (2) (up to 1.0 M) with recombinant Escherichia coli overproducing a versatile carbonyl reductase called SCR (Saccharomyces cerevisiae carbonyl reductase) together with a glucose dehydrogenase (GDH).2 A remarkable temperature effect on productivity was observed in the whole-cell reduction of 2, and the optimum productivity as high as 178 g L1 was attained at 20 °C (Scheme 9.1). Cl

CO2Me N S clopidogrel

Cl

Cl

O

OH CO2Me

CO2Me 2

SCR NADPH

gluconolactone gluconic acid

(R)-1 NADP+

GDH

glucose

recombinant E. coli

Scheme 9.1

9.6.1 9.6.1.1 • • • • • •

Procedure 1: Cultivation of Recombinant E. coli Materials and Equipment

Ampicillin (250 mg) chloramphenicol (85 mg) E. coli BL21(DE3) cells harboring pESCR and pABGD Luria–Bertani (LB) medium: tryptone (25 g), yeast extract (13 g), NaCl (25 g) isopropyl--D-thiogalactopyranoside (IPTG, 60 mg) Milli-Q water (2.5 L)

292

• • • • • • • • • •


0.1 M phosphate buffer (200 mL) test tube ( 8) rotary shaker sterile toothpicks 1 L Erlenmeyer flask ( 8) measuring cylinder cotton plug autoclave UV–vis spectrophotometer centrifuge.

9.6.1.2

Procedure

1. E. coli BL21(DE3) cells harboring pESCR and pABGD, previously constructed,3 were grown in LB medium (3 mL 8) containing ampicillin (100 mg mL1) and chloramphenicol (34 mg mL1) at 37 °C for 15 h with shaking at 230 rpm. 2. The culture (3 mL 8) was transferred to the same medium (300 mL 8) in a 1 L Erlenmeyer flask and shaken at 200 rpm at 37 °C. 3. IPTG (0.1 mM) was added when optical density at 600 nm reached 0.6–0.8. The cells were further incubated at 28 °C for 18 h with shaking at 200 rpm and then harvested by centrifugation (7000 rpm, 4 °C, 10 min) into four portions. 4. Each of the four portions was washed with 0.1 M phosphate buffer (pH 7.0, 50 mL). 5. The wet cell pellet (7–8 g) obtained was stored at 20 °C until it was used for asymmetric reduction. 9.6.2

Procedure 2: Synthesis of Methyl (R)-o-Chloromandelate ((R)-1) Cl

Cl

O

OH

recombinant E. coli

CO2Me

CO2Me

NADP+, glucose, buffer 2

9.6.2.1 • • • • • • • • • • •

(R)-1


Cells prepared above (2 g) cell pellet of recombinant E. coli (2.0 g) methyl o-chlorobenzoylformate (2) (1.98 g) D-glucose (3.6 g) nicotinamide adenine dinucleotide phosphate (NADPþ, 10 mg) 0.1 M phosphate buffer (pH 7.0, 10 mL) 2 M NaOH (5 mL) NaCl (5.5 g) MgSO4 (1 g) silica gel hexane

9.6 Enantioselective and Efficient Synthesis of Methyl (R)-o-Chloromandelate

• • • • • • • • • • •

293

ethyl acetate 100 mL test tube (2.7 cm diameter) water bath with a thermostat magnetic stirrer vortex mixer pH indicator paper centrifuge rotary evaporator 200 mL Erlenmeyer flask 200 mL round-bottom flask 30 mL round-bottom flask.

9.6.2.2

Procedure

1. To a mixture of glucose (3.60 g, 20.0 mmol), NADPþ (10 mg, 12 mmol), and E. coli BL21(DE3) cells harboring pESCR and pABGD (2.0 g) in 0.1 M phosphate buffer (pH 7.0, 10 mL) in a 100 mL test tube was added methyl o-chlorobenzoylformate (2) (1.98 g, 10.0 mmol). 2. The mixture was stirred in a water bath at 20 °C for 24 h, during which 2 M NaOH was added to maintain pH 7 by neutralizing the acid formed in the progress of the reaction. 3. Solid NaCl (5.5 g) was added and the product was extracted with EtOAc (25 mL 3). Phase separation was effected by centrifugation (3200 rpm, 10 min). The combined organic layers were dried over MgSO4, filtered and concentrated under reduced pressure. Purification by silica-gel column chromatography (hexane/ EtOAc (10:1)) gave methyl (R)-o-chloromandelate ((R)-1) as a colorless oil (1.78 g, 89 %). [] 19 D ¼ 178.3 (c ¼ 1.3, CHCl3), >99 % ee, (R). High-performance liquid chromatography (HPLC): Chiralpak AD-H (Daicel Chemical Industries, Ltd), hexane/i-PrOH (9:1), flow rate 0.5 mL min1, detection 254 nm, (S) 20.3 min, (R) 22.7 min. 1H NMR (CDCl3, 600 MHz) 3.56 (d, J ¼ 5.4 Hz, 1H), 3.78 (s, 3H), 5.57 (d, J ¼ 5.4 Hz, 1H), 7.28–7.29 (m, 2H), 7.39–7.40 (m, 2H). 13C NMR (CDCl3, 150 MHz) 53.2, 70.3, 127.2, 128.8, 129.8, 130.0, 133.5, 135.9, 173.7. IR (film) 3454, 3003, 2955, 1744, 1441, 1223, 1090, 756 cm1. 9.6.3

Conclusion

An efficient and green chemoenzymatic method for methyl (R)-o-chloromandelate ((R)-1) has been developed. The asymmetric reduction of methyl o-chlorobenzoylformate (2) with recombinant E. coli overproducing a versatile carbonyl reductase, SCR, gave (R)-1 with >99% ee. This is the first example of the direct asymmetric synthesis of (R)-1 with >99 % ee. A remarkable temperature effect on productivity was observed in the whole-cell reduction of 2, and the optimum productivity as high as 178 g L1 was attained at 20 °C (Table 9.4). The bioreduction of 2 is a green process, because the hydride source is glucose, which is a cheap biomass-derived reagent, and because the E. coli catalyst can be multiplied easily and inexpensively. Moreover, the bioreduction is performed in an aqueous solution under air.

294


Table 9.4

Asymmetric reduction of 2 with recombinant E. coli.a

Entry

[2] (M)

[2] (g L1)

T (°C)

0.3 0.3 0.6 1.0 1.0 1.0

60 60 120 198 198 198

30 25 25 25 20 15

1 2 3 4 5 6

C (%)b 92 >99 94 90 99 86

Yield (%)c

Ee (%)d

76 88 88 85 89 82

>99 >99 >99 >99 >99 >99

a

Conditions: 2 (0.60–1.98 g, 3.0–10.0 mmol), wet cells of E. coli BL21(DE3) harboring pESCR and pABGD (2.0 g), glucose (2 equiv), NADPþ (10 mg, 12 mmol), 0.1 M phosphate buffer (pH 7.0, 10 mL). Conversion determined by 1H NMR. c Isolated yield of (R)-1. d Determined by HPLC (Chiralpak AD-H, hexane/i-PrOH (9:1)). b

References 1. Grimley, J., Pharma challenged. Chem. Eng. News, 2006, Dec. 4, 17–28. 2. Ema, T., Okita, N., Ide, S., Sakai, T., Highly enantioselective and efficient synthesis of methyl (R)-o-chloromandelate with recombinant E. coli: toward practical and green access to clopidogrel. Org. Biomol. Chem., 2007, 5, 1175–1176. 3. Ema, T., Yagasaki, H., Okita, N., Takeda, M., Sakai, T., Asymmetric reduction of ketones using recombinant E. coli cells that produce a versatile carbonyl reductase with high enantioselectivity and broad substrate specificity. Tetrahedron, 2006, 62, 6143–6149.

10 Reduction of Functional Groups 10.1

Reduction of Carboxylic Acids by Carboxylic Acid Reductase Heterologously Expressed in Escherichia coli Andrew S. Lamm, Arshdeep Khare and John P.N. Rosazza*

The biocatalytic reduction of carboxylic acids to their respective aldehydes or alcohols is a relatively new biocatalytic process with the potential to replace conventional chemical processes that use toxic metal catalysts and noxious reagents. An enzyme known as carboxylic acid reductase (Car) from Nocardia sp. NRRL 5646 was cloned into Escherichia coli BL21(DE3).1–7 This E. coli based biocatalyst grows faster, expresses Car, and produces fewer side products than Nocardia. Although the enzyme itself can be used in small-scale reactions, whole E. coli cells containing Car and the natural cofactors ATP and NADPH, Hþ are easily used to reduce a wide range of carboxylic acids, conceivably at any scale. The biocatalytic reduction of vanillic acid to the commercially valuable product vanillin is used to illustrate the ease and efficiency of the recombinant Car E. coli reduction system.4 A comprehensive overview is given in Reference 6, and experimental details below are taken primarily from Reference 7. 10.1.1

Biocatalytic Synthesis of Vanillin O

O

OH

Aldehyde reductase

Car, ATP, NADPH OMe

Mg+2

OH Vanillic acid


HO

H

OMe

OMe OH

OH

Vanillin

Vanillyl Alcohol


296

Reduction of Functional Groups

10.1.1.1

Materials and equipment

• E. coli BL21(DE3) harboring plasmid pPV2.85 (frozen glycerol stocks) • Luria–Bertani (LB) broth powder (20 g L1) • LB agar powder (15 g L1) • ampicillin (100 mg mL1 stock solution in water, filter sterilized) • high-performance liquid chromatography (HPLC)-grade acetonitrile (800 mL) • HPLC-grade water (200 mL) • HPLC-grade formic acid (1 mL) • sodium dihydrogen phosphate (12g.L-1) • sodium hydrogen carbonate (5 g) • sodium vanillate stock solution (50 mg mL1) • vanillyl alcohol (1 g as HPLC or thin-layer chromatography (TLC) standard) • vanillin (1 g as HPLC or TLC standard) • 0.22 mm polyvinylidene difluoride syringe filters • 10 mL and 1 mL syringes • sterile loop • Petri dish • two 125 ml and two 1 L stainless-steel-capped DeLong flasks • rotary shakers at 37 C • centrifuge capable of reaching 5000g while holding 4 C • HPLC system and UV detection • Econosil HPLC column (C18, 5 mm, 150 mm 3.2 mm; Alltech). Optional • • • • • • • • •

Silica gel TLC plates (silica gel 60 F254, Merck) 30 % w/v phosphomolybdic acid in ethanol (100 mL) reagent spray bottle heat gun UV lamp/viewing box benzoic acid (50 mg mL1 in water) 3-chlorobenzoic acid (50 mg mL1 in water) 4-chlorobenzoic acid (50 mg mL1 in water) 3-(4-hydroxy-3-methoxyphenyl)-propenoic acid (50 mg mL1 in water).

10.1.1.2

Procedure

Initial culture 1. Crystals from a frozen glycerol stock of E. coli BL21(DE3)/pPV2.85 were streaked onto LB agar plates with ampicillin (100 mg mL1) to obtain single colonies. 2. Single colonies were inoculated into 20 mL of LB medium (containing 100 mg mL1 ampicillin) in 125 mL stainless-steel-capped DeLong flasks. 3. Cultures were incubated with shaking at 250 rpm on a rotary shaker at 37 C. A 1 % inoculum derived from 8 h stage I cultures was used to initiate fresh LB cultures (200 mL) with antibiotics in a 1 L DeLong flask. These cultures were incubated at 37 C for 16 h with shaking at 250 rpm. 4. Car-containing E. coli cells were pelleted by centrifugation at 5000g for 6 min at 4 C.

10.1 Reduction of Carboxylic Acids by Carboxylic Acid Reductase

297

Whole-cell Carboxylic Acid Reduction 1. Car-containing E. coli cells were resuspended in 200 mL of 0.9 % (w/v) NaCl and pelleted once again by centrifugation at 5000g for 6 min at 4 C. 2. A sodium vanillate stock solution (50 mg mL1) was prepared by dissolving equimolar amounts of vanillic acid and NaHCO3 in 0.1 M Na2HPO4 (pH 7). Reaction mixtures of 50 mL contained 0.4 % glucose, 1.5 g of wet E. coli cells, 200 mg of sodium vanillate in pH 7, 0.1 M Na2HPO4. 3. Reactions were incubated at 28 C with shaking at 220 rpm and 1 mL samples were withdrawn at various time intervals for analysis. Samples are treated as described in Section 10.1.1.3. 10.1.1.3

Analytical Methods

Standard solutions were prepared by dissolving weighed amounts of compounds in a 1:1 (v/v) mixture of pH 7, 0.1 M Na2HPO4/acetonitrile. Aliquots of 0.5 mL of biotransformation samples were mixed with 0.5 mL of acetonitrile and mixtures were vortexed for 30 s. After standing at room temperature for 30 min, samples were microcentrifuged at 20 000g for 3 min, the supernatants filtered through 0.22 mm polyvinylidene difluoride syringe filters and 1–2 ml injected for HPLC analysis. The HPLC system used a mobile phase consisting of CH3CN/H2O/HCOOH (20:80:1, v/ v/v). Quantitation of standards and samples was achieved by isocratic elution over a C18, 5 mm, Econosil HPLC column at a flow rate of 0.4 mL min1. HPLC retention volumes and detection wavelengths for standards were as follows: vanillyl alcohol, 1.7 mL and 277 nm; vanillic acid, 2.5 mL and 260 nm; and vanillin, 4.1 mL and 284 nm. TLC analysis of samples was conducted on silica-gel plates carefully spotted with 10– 20 mg of standard compounds, and 30 mL of bioconversion reaction samples. Plates developed with 75:25:1 (v/v/v) CH2Cl2/CH3CN/HCOOH solvent may be visualized with a 254 nm UV lamp and/or by spraying with a 30 % w/v phosphomolybdic acid/ 95% ethanol spray reagent followed by gentle heating. Rf values of standards are: vanillyl alcohol, 0.8; vanillic acid, 0.5; and vanillin 0.4. Other suitable alternate aromatic carboxylic substrates include 3-chlorobenzoic acid, 4chlorobenzoic and 3-(4-hydroxy-3-methoxyphenyl)-propenoic acid. 10.1.2

Conclusion

Most of the vanillic acid was reduced by E. coli containing Car in 2 h to vanillin (80 %) and vanillyl alcohol (20 %). Car does not reduce aldehydes to alcohols. However, E. coli’s endogenous aldehyde reductase/dehydrogenase reduces vanillin to vanillyl alcohol. The broad substrate specificity of Car enables the wide application of this biocatalyst to other important applications, such as enantiomeric resolution of isomers such as ibuprofen1 and the reductions of many other natural and synthetic carboxylic acids.

References 1. Chen, Y. and Rosazza, J.P.N., Microbial transformation of ibuprofen by a Nocardia species. Appl. Environ. Microbiol., 1994, 60, 1292–1296. 2. Li, T. and Rosazza, J.P.N. Purification, characterization, and properties of an aryl aldehyde oxidoreductase from Nocardia sp. strain NRRL 5646. J. Bacteriol., 1997, 179, 3482–3487.

298


3. Li, T. and Rosazza, J.P.N., NMR Identification of an acyl-adenylate intermediate in the arylaldehyde oxidoreductase catalyzed reaction. J. Biol. Chem., 1998, 273, 34230–34233. 4. Li, T. and Rosazza, J.P.N., Biocatalytic synthesis of vanillin. Appl. Environ. Microbiol., 2000, 66, 684–687. 5. He, A., Li, T., Daniels, L., Fotheringham, I. and Rosazza, J.P.N. Nocardia sp. carboxylic acid reductase: cloning, expression, and characterization of a new aldehyde oxidoreductase family. Appl. Environ. Microbiol., 1994, 70, 1874–1881. 6. Venkitasubramanian, P., Daniels, L. and Rosazza, J.P.N., Biocatalytic reduction of carboxylic acids: mechanism and application. In Biocatalysis in the Pharmaceutical and Biotechnology Industries, Patel, R. (ed). CRC Press LLC: Boca Raton, FL, 2006, pp. 425–440. 7. Venkitasubramanian, P., Daniels, L. and Rosazza, J.P.N., Reduction of carboxylic acids by Nocardia aldehyde oxidoreductase requires a phosphopantetheinylated enzyme. J. Biol. Chem., 2007, 282, 478–485.

10.2 Light-driven Stereoselective Biocatalytic Oxidations and Reductions

10.2

299

Light-driven Stereoselective Biocatalytic Oxidations and Reductions Andreas Taglieber, Frank Schulz, Frank Hollmann, Monika Rusek and Manfred T. Reetz*

Recently, the use of visible light to promote the direct reductive regeneration for flavindependent enzymes has been proposed.1 The feasibility of this concept has been successfully demonstrated for stereoselective Baeyer–Villiger oxidations using an engineered variant of phenylacetone monooxygenase (PAMO-P31–3) from Thermobifida fusca and for the enantioselective reduction of ketoisophorone catalyzed by YqjM from Bacillus subtilis.4 The light-driven regeneration system is based on the use of flavin cocatalysts that are activated by white light and react from their excited state with a sacrificial electron donor (ethylenediaminetetraacetic acid (EDTA)). In this reaction, the flavin cocatalyst is converted into a reduced species which transfers the reducing equivalents to the enzymebound flavin, thereby regenerating the reduced enzyme (Scheme 10.1). O2 A

EDTA

flavin ox light

O

E-FADred R

R'

BVMO O

decomposition products

flavinred

E-FADox H2O

R

O

R'

O R2 B

EDTA

flavin ox light

decomposition products

E-FMNred

R1

R3

R4

YqjM

O H

flavinred

E-FMNox

R1

R2 R3

H

R4

Scheme 10.1 Light-driven regeneration of (A) a Baeyer–Villiger monooxygenase (BVMO) and (B) for the flavin-dependent reductase YqjM

By means of this reaction, the use of the costly and unstable natural redox cofactor reduced nicotinamide adenine dinucleotide phosphate (NADPH) was circumvented and the reactions were carried out in a straightforward procedure in a chemical laboratory (Scheme 10.2, Table 10.1). O

O light, YqjM, FMN, EDTA

100% conv. 83% ee

4 h, 30 °C O 6

Scheme 10.2

O (R)-7

Light-driven YqjM-catalyzed reduction of ketoisophorone

300

Reduction of Functional Groups Table 10.1

Light-driven PAMO-P3-catalyzed Baeyer–Villiger oxidations. O

O R

light, PAMO-P3 FAD, EDTA 30 °C

r ac-1 a R = C 6H 5 b R = CH 2 C6 H5

O

10.2.1.1 • • • • • • • • • • • • • •

R R

+ (S)- 1

(R)-2

light, PAMO-P3 FAD, EDTA 30 °C

rac -3 Substrate 1a 1b 3

10.2.1

O O

(–)- 4 Conversion(%) 48 30 93

O O O

+

O (–)-5 Ee(%) of product 97 97 4: 92; 5: ≥95

Procedure 1: Expression and Purification of PAMO-P3 Materials and equipment

Terrific broth5 carbenicillin L-arabinose 5 L fermenter (Labfors HT, Infors) centrifuge sonicator flavine adenine dinucleotide (FAD) lysozyme (from chicken egg-white) Fractogel His-Bind resin (Novagen) 500 mL glass column (with glass frit at the bottom) imidazole centrifuge filters (Amicon, molecular weight cutoff (MWCO) 10 000) PD-10 desalting columns (GE Healthcare) Escherichia coli TOP10 [pPAMO-P3].3

10.2.1.2

Procedure

1. For the expression of PAMO-P3, 100 mL of an overnight preculture of E. coli TOP10 [pPAMO-P3] in terrific broth medium supplemented with 100 mg L1 of carbenicillin was used to inoculate 5 L terrific broth medium supplemented with 100 mg L1 of carbenicillin and 0.1 % L-arabinose in a 5 L fermenter. The expression was carried out


2.

3.

4.

5.

6.

301

over the course of 7 h at 37 C and constant stirrer revolution (800 rpm). The cells were harvested by centrifugation (10 000g, 4 C, 15 min) and frozen at 80 C. After thawing, the complete cell paste was suspended in 250 mL of 20 mM potassium phosphate buffer (pH 7.4) containing 10 mM of FAD and 0.2 mg mL1 of lysozyme and incubated at 4 C for 30 min. Cells were disrupted by sonication and the lysate clarified by centrifugation (10 000g, 4 C, 60 min). The supernatant was incubated at 50 C for 1 h in a water bath and subsequently centrifuged. The supernatant was supplemented with NaCl to a concentration of 0.5 M NaCl and mixed with 35 mL of Novagen Fractogel His-Bind resin (pre-equilibrated and loaded with Ni2þ as recommended by the manufacturer). The suspension was gently mixed for 30 min and then manually loaded into a 500 mL glass column and packed under 1.5 bar Ar pressure. The material eluted was loaded onto the column once more; subsequently, the column was washed with 350 mL of 20 mM KH2PO4 (pH 7.4) and then with 350 mL of 20 mM potassium phosphate buffer (pH 7.4) supplemented with 1 mM imidazole. PAMO-P3 was eluted with 100 mL of 50 mM tris-HCl (pH 7.4) containing 200 mM imidazole. In total, 25 mL of yellow eluate was collected and concentrated to 12.5 mL by centrifuge filters (Amicon, MWCO 10 000). The final purification step in each case was desalting of the eluates via PD-10 columns (Amersham, 8.3 mL Sephadex G-25 medium) according to the recommendations of the column manufacturer, using 50 mM tris-HCl (pH 7.4) as equilibration and elution buffer. The concentration of purified enzyme in 50 mM tris-HCl (pH 7.4) was determined by the UV–vis absorbance at 441 nm ("441 nm ¼ 12.4 mM1 cm1).6

10.2.2 10.2.2.1

Procedure 2: Light-driven PAMO-P3-catalyzed Baeyer–Villiger Oxidations Materials and Equipment

• PAMO-P3, purified enzyme • NADPþ • substrate (2-phenylcyclohexanone, 2-benzylcyclohexanone or bicyclo[3.2.0]hept-2-en6-one) • EDTA • flavin cocatalyst (FAD, flavin mononucleotide (FMN) or riboflavin) • 100 W white-light bulb. 10.2.2.2

Procedure

1. A final reaction volume of 250 mL, containing 10 mM PAMO-P3, 25 mM EDTA, 100 mM FAD, 250 mM NADPþ, 1 mM or 2 mM substrate and 50 mM tris-HCl (pH 7.4), was incubated under aerobic conditions at 30 C in a water bath exposed to light from a 100 W Osram white-light bulb. The light was filtered through 1 cm of water and 0.5 cm of DURANTM glass. The approximate distance between light source and the reaction vessels was 6 cm. The reaction mixture was extracted with 275 mL of ethyl acetate each and analyzed by gas chromatography (GC). The reaction took around 6 h and was followed by GC analysis using authentic standards.

302


For all experiments, PAMO-P3 was produced and purified as described above to allow for accurate quantification of the results. However, the reaction also works using crude enzyme as obtained after bacterial lysis.2 10.2.2.3

Characterization of the Products

For all products, synthetic standards were prepared by meta-chloroperbenzoic acidmediated Baeyer–Villiger oxidation.7 The crude products were purified by silica-gel chromatography and the NMR spectra matched the data reported in the literature (see below). Stereochemical configurations were assigned based on analogous conversions carried out using cyclohexanone monooxygenase. All GC results were confirmed by GC– mass spectrometry (MS) analysis (instrument: Finnigan SSQ7000; GC–electron impact (EI), achiral GC methods described below). 10.2.2.4

GC Analyses

Compound 2a8 Achiral method. Instrument: Agilent Technologies 6890N; carrier gas: 0.6 bar H2; column: 15 m ZB1 (100 % dimethylpolysiloxane, 0.25 mm inner diameter, 0.5 mm film); injector T ¼ 220 C, detector T ¼ 350 C; program: ramp 80 C to 195 C with 8 C min1, then 20 C min1 to 340 C; retention times: 7.96 min (1a), 10.85 min (2a). GC-factor correction was performed versus n-C16 standard; correction factor: 1.15. Chiral method. Instrument: Agilent Technologies 6890N; carrier gas: 0.6 bar H2; column: 30 m BGB-176 (20 % 2,3-dimethyl-6-tert-butyldimethylsilyl--cyclodextrin dissolved in BGB-15, 0.25 mm inner diameter, 0.1 mm film); injector T ¼ 220 C, detector T ¼ 350 C; program: 150 C (iso) 10.5 min, ramp to 160 C with 50 C min1, 160 C (iso) 16 min; retention times: (S)-1a, 9.56 min; (R)-1a, 9.77 min; (S)-2a, 21.20 min; (R)-2a, 21.51 min. Compound 2b8 Achiral method. Instrument: Agilent Technologies 6890N; carrier gas: 0.6 bar H2; column: 15 m ZB1 (100 % dimethylpolysiloxane, 0.25 mm inner diameter, 0.5 mm film); injector T ¼ 220 C, detector T ¼ 350 C; program: ramp 80 C to 320 C with 8 C min1, then 320 C (iso) 1 min; retention times: 9.59 min (1b), 12.46 min (2b). Chiral method. Instrument: Agilent Technologies 6890N; carrier gas: 0.6 bar H2; column: 30 m BGB-176 (20 % 2,3-dimethyl-6-tert-butyldimethylsilyl--cyclodextrin dissolved in BGB-15, 0.25 mm inner diameter, 0.1 mm film); injector T ¼ 220 C, detector T ¼ 350 C, program: 150 C (iso) 12.5 min, ramp to 200 C with 100 C min1, 200 C (iso) 5 min; retention times: (R)-1b, 11.97 min; (S)-1b, 11.72 min; (R)-2b, 17.12 min; (S)-2b, 16.86 min. Compounds 4 and 59,10 Achiral method. Instrument: Agilent Technologies 6890N; carrier gas: 0.6 bar H2; column: 30 m RTX-5 (5 % diphenylpolysiloxane, 95 % dimethylpolysiloxane, 0.25 mm inner diameter, 0.25 mm film); injector T ¼ 220 C, detector T ¼ 350 C; program: ramp 60 C to 330 C with 6 C min1, then 350 C (iso) 10 min, retention times: 4.60 min (3), 9.79 min (4), 9.91 min (5).


303

Chiral method. Agilent Technologies 6890N; carrier gas: 0.7 bar H2; column: 30 m BGB-178 (20 % 2,3-diethyl-6-tert-butyldimethylsilyl--cyclodextrin dissolved in BGB-15, 0.25 mm inner diameter, 0.25 mm film); injector T ¼ 220 C, detector T ¼ 350 C; program: 125 C (iso) 14.5 min, ramp to 230 C with 10 C min-1, 230 C (iso) 5 min; retention times: 10.61 min (4), 11.30 min (þ5), 11.98 min (5), 12.50 min (þ4). 10.2.3 10.2.3.1 • • • • • • • • •

Procedure 3: Expression of YqjM Materials and Equipment

IPTG E. coli Rosetta (DE3) [pET21a-YqjM]11 terrific broth medium5 FMN carbenicillin 5 L fermenter (Labfors HT, Infors) centrifuge sonicator PD-10 desalting columns (GE Healthcare).

10.2.3.2

Procedure

Expression of YqjM was carried out in a 5 L fermenter using terrific broth medium supplemented with 100 mg L1 carbenicillin. As an inoculum, 100 mL of preculture was used. Temperature and stirrer speed were kept constant (800 rpm, 37 C). At an optical density at 600 nm of 0.67, 500 mL of 1 M IPTG was added and the temperature adjusted to 30 C. After 4 h of expression, the cells were harvested by centrifugation and the resulting cell paste frozen at 80 C overnight. After thawing, the complete cell paste was suspended in 100 mL of 50 mM tris-HCl buffer (pH 7.4). Cell lysis was achieved by sonication. DNAse I (0.1 mg mL1) was added and the crude lysate incubated at 4 C for 30 min. The lysate was clarified by centrifugation (10 000g, 4 C, 60 min). After the reconstitution of the enzyme with FMN (1 h incubation at 4 C in the presence of 100 mM FMN) followed by removal of excess FMN using PD-10 desalting columns (GE Healthcare) according to the recommendations of the column manufacturer with 50 mM tris-HCl (pH 7.4) as equilibration and elution buffer, the protein was analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis using a 12.5 % gel. Using the densitograph function of BioDocAnalyze (Biometra), the YqjM content was determined to be 32 % of the total protein content of the protein preparation. The total protein content was determined using a Bradford assay reagent (Bio-Rad) with bovine serum albumin as standard.12 10.2.4 10.2.4.1

Procedure 4: Light-driven YqjM-catalyzed Reduction of Ketoisophorone Materials and equipment

• YqjM • ketoisphorone

304


• EDTA • FMN • 100 W white-light bulb 10.2.4.2

Procedure

1. A final reaction volume of 250 mL, containing 2.6 mM YqjM (corresponds to approximately 0.1 mg mL1 total protein content), 25 mM EDTA, 100 mM FMN, 1 mM ketoisophorone and 50 mM tris-HCl (pH 7.4), was incubated under anaerobic conditions (closed reaction vessel with small head space) at 30 C in a water bath exposed to the light of a 100 W Osram white-light bulb for 4 h. The experimental setup was the same as described in Procedure 2 (Section 10.2.2). 2. Reactions were followed and analyzed by GC. For GC analysis, samples were extracted with an equal volume of ethyl acetate and analyzed. Peak assignment was performed with an authentic standard of the product and confirmed by GC–MS (instrument: Finnigan SSQ7000, GC–EI, achiral GC method described below). 10.2.4.3

GC-Analyses

Achiral method. Instrument: Agilent Technologies 6890N; carrier gas: 0.6 bar H2; column: 15 m ZB1 (100 % dimethylpolysiloxane, 0.25 mm inner diameter, 0.5 mm film); injector T ¼ 220 C, detector T ¼ 350 C; program: ramp 80 C to 110 C with 5 C min1, then 20 C min-1 to 340 C; retention times: 2.83 min (6), 3.05 min (7). Chiral method. Agilent Technologies 6890N; carrier gas: 0.6 bar H2; column: 30 m BGB176 (20 % 2,3-dimethyl-6-tert-butyldimethylsilyl--cyclodextrin dissolved in BGB15, 0.25 mm inner diameter, 0.1 mm film); injector T ¼ 220 C, detector T ¼ 350 C; program: 100 C (iso) 15 min; retention times: 12.11 min (6); 12.35 min ((R)-7) and 13.90 min ((S)-7). 10.2.5

Conclusion

The straightforward concept for the direct light-driven regeneration of flavin-dependent enzymes has been successfully applied for two representative classes of such enzymes: a reductase and a monooxygenase. Therefore, it can be expected that this concept can also be applied to other flavin-dependent enzymes, potentially leading to additional practical catalyst systems for applications in synthetic organic chemistry.

References 1. Hollmann, F., Taglieber, A., Schulz, F. and Reetz, M.T., A light-driven stereoselective biocatalytic oxidation. Angew. Chem. Int. Ed., 2007, 46, 2903. 2. Schulz, F., Leca, F., Hollmann, F. and Reetz, M.T., Towards practical biocatalytic Baeyer– Villiger reactions: applying a thermostable enzyme in the gram-scale synthesis of opticallyactive lactones in a two-liquid-phase system. Beilstein J. Org. Chem., 2005, 1, 10. 3. Bocola, M., Schulz, F., Leca, F., Vogel, A., Fraaije, M.W. and Reetz, M.T., Converting phenylacetone monooxygenase into phenylcyclohexanone monooxygenase by rational design: towards practical Baeyer–Villiger monooxygenases. Adv. Synth. Catal., 2005, 347, 979.


305

4. Taglieber, A., Schulz, F., Hollmann, F., Rusek, M. and Reetz, M.T., Light-driven biocatalytic oxidation and reduction reactions: scope and limitations. ChemBioChem, 2008, 9, 565. 5. Sambrook, J. and Russel, D., Molecular Cloning: A Laboratory Manual, 3rd edn. Cold Spring Harbor Laboratory Press, New York, 2000. 6. Fraaije, M.W., Wu, J., Heuts, D.P.H.M., van Hellemond, E.W., Spelberg, J.H.L. and Janssen, D.B., Discovery of a thermostable Baeyer–Villiger monooxygenase by genome mining. Appl. Microbiol. Biotechnol. 2005, 66, 393. 7. Krow, G.R., The Baeyer–Villiger oxidation of ketones and aldehydes. In Organic Reactions, vol. 43, Paquette, L.A. (ed.). John Wiley & Sons, Inc.: New York, 1993, pp. 251–798. 8. Alphand, V., Furstoss, R., Pedragosa-Moreau, S., Roberts, S.M. and Willetts, A.J., Comparison of microbiologically and enzymatically mediated Baeyer–Villiger oxidations: synthesis of optically active caprolactones. J. Chem. Soc. Perkin Trans. 1, 1996, 1867. 9. Hudlicky, T., Reddy, D.B., Govindan, S.V., Kulp, T., Still, B. and Sheth, J.P., Intramolecular cyclopentene annulation. 3. Synthesis and carbon-13 nuclear magnetic resonance spectroscopy of bicyclic cyclopentene lactones as potential perhydroazulene and/or monoterpene synthons. J. Org. Chem., 1983, 48, 3422. 10. Grieco, P.A., Cyclopentenones. Efficient synthesis of cis-jasmone. J. Org. Chem., 1972, 37, 2363. 11. Fitzpatrick, T.B., Amrhein, N. and Macheroux, P., Characterization of YqjM, an old yellow enzyme homolog from Bacillus subtilis involved in the oxidative stress response. J. Biol. Chem., 2003, 278, 19891. 12. Bradford, M.M., A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Anal. Biochem., 1976, 72, 248.

306


10.3

Unnatural Amino Acids by Enzymatic Transamination: Synthesis of Glutamic Acid Analogues with Aspartate Aminotransferase Thierry Gefflaut*, Emmanuelle Sagot and Jean Bolte

Aminotransferases (ATs) catalyse the stereoselective transfer of the amino group from a donor substrate to an acceptor prochiral carbonyl derivative. ATs are very common enzymes with high specific activities and relaxed substrate specificity. The development of equilibrium shifted transamination processes allowed the preparation of a variety of biologically active compounds, including unnatural L- and D--amino acids1,2 as well as aminoacids or simple amines.3–6 Aspartate aminotransferase (AspAT) offers the opportunity to shift the transamination equilibrium through the use of cysteine sulfinic acid (CSA) as the amino donor substrate: CSA, which is a close analogue of aspartic acid, is converted into the very unstable pyruvyl sulfinic acid, which spontaneously decomposes into pyruvic acid and sulfur dioxide, thus providing the equilibrium shift. Recently, AspAT has proven useful for the stereoselective preparation of a variety of neuroactive glutamic acid derivatives.7–10 This methodology is exemplified below with the preparation of (2S,4R)-4methyl Glu (2), a potent selective ligand for kainate receptors: AspAT gives exclusively the L-amino acid and allows the kinetic resolution of the racemic -keto acid substrate 1 readily prepared by conventional chemical methods.7 This catalyst thus offers the stereocontrol of two asymmetric centres. 10.3.1

Procedure 1: Synthesis of (2S,4R)-4-Methyl Glutamic Acid AspAT O

HO

OH O

HO

O

+

HO

O

(2S,4R)-2 NH 2 O S

HO O

• • • • • • • • • • •

OH O

rac-1

10.3.1.1

O

NH 2

H2O, pH7.6

O OH

CSA

+ SO2

HO O


4-Methyl-2-oxoglutaric acid (1)7 (0.5 g, 2.9 mmol) CSA (0.45 g, 2.9 mmol) acetaldehyde (128 mg, 2.9 mmol) AspAT (from pig heart, Sigma) (1 mg) KH2PO4 (1.36 g, 10 mmol) KOH lactate dehydrogenase (from rabbit muscle, Sigma) (0.1 mg mL1) reduced nicotinamide adenine dinucleotide (NADH, 10 mg mL1) Dowex 50WX8 (200–400 mesh, Hþ form) (20 g) Dowex 1X8 (200–400 mesh, AcO form) (20 g) 1 M NH4OH (100 mL)

OH O

O

10.3 Unnatural Amino Acids by Enzymatic Transamination

• • • • • • • • • • • • • • •

307

1 M AcOH (200 mL) ninhydrin (0.2 g in 100 mL EtOH) (for thin-layer chromatography (TLC) dip) propan-1-ol (50 mL) (for TLC elution) pH meter 250 mL flask magnetic stirrer 1 mL adjustable volume pipette 20 mL adjustable volume pipette 1.5 mL microcentrifuge tubes (for enzyme solutions) centrifuge (for microtubes) 1.5 mL disposable cuvettes (for spectrophotometry at 340 nm) UV or visible spectrophotometer equipment for column chromatography (column: 2 cm 20 cm, tubes 5–10 mL) TLC plates (silica gel 60F254, Merck) rotatory evaporator.

10.3.1.2

Procedure

1. In a 250 mL flask equipped with a stir bar were introduced 4-methyl-2-oxoglutaric acid 1 (0.5 g, 2.9 mmol), CSA (0.45 g, 2.9 mmol), water (145 mL) and acetaldehyde (128 mg, 2.9 mmol).11 The pH of the solution was adjusted to 7.6 with 1 M KOH before the addition of pig heart AspAT (1 mg). The commercial enzyme suspension in 3 M (NH4)2SO4 was centrifuged (5 min at 10 000 rpm), the supernatant eliminated and the enzyme pellet dissolved in the reaction mixture. The reaction was stirred slowly at room temperature and monitored by titration of pyruvic acid formed from CSA. 2. The solutions needed for pyruvic acid titration were all prepared in 0.1 M potassium phosphate buffer pH 7.6. In a disposable 1.5 mL cuvette were introduced a 10 mg mL-1 solution of NADH (20 ml), a 0.1 mg mL1 solution of rabbit muscle lactate dehydrogenase (10 mL, 1.2 unit) and phosphate buffer (965 mL). The initial optical density (ODi £ 1.5) was measured at 340 nm. An aliquot of the reaction mixture (5 mL) was then added and the final stable OD (ODf) was measured. Pyruvic acid concentration in the reaction mixture was calculated using "NADH ¼ 6220 M1 cm1: [Pyruvate] ¼ (ODi ODf) 200/6220. 3. When a conversion of 40 % was reached (8 mM pyruvate formed in 2–3 h), the reaction mixture was rapidly passed through a column of Dowex 50WX8 resin (Hþ form, 2 cm 10 cm). The column was then washed with water (100 mL) until complete elution of residual substrate 1, pyruvic acid and CSA. It was then eluted with 1 M NH4OH (100 mL). The fractions (5 mL) were analysed by TLC (eluent n-PrOH/H2O, 7:3, v/v). The ninhydrinpositive fractions were combined and concentrated to dryness under reduced pressure. The purity of the product was further increased by anion-exchange chromatography: 4. The residue was diluted in water (5 mL) and, if necessary, the pH adjusted to 7.0 with 1 M KOH before adsorption of the product on a column of Dowex 1X8 resin (200–400 mesh, AcO form, 2 cm 10 cm). The column was washed with water (50 mL) and then eluted with AcOH aqueous solutions (50 mL of 0.1 M, 50 mL of 0.2 M and 50 mL of 0.5 M AcOH). The ninhydrin-positive fractions were combined and dried under reduced pressure to afford (2S,4R)-4-methyl glutamic acid 2 isolated as a white solid (192 mg, 41 %) and with a high purity (>98 %).

308

Reduction of Functional Groups H2 N

H2N

Alk

HO

OH O

OH O

O

H2N

O

O

O H2N

O H2N

n

O

O

HO

OH

HO

OH

HO O

HO

Alk = Me, Et, Pr, Bu, Pn iPr, iBu, iPn, Bn

OR

H2N

HO

O

H2N n = 1,2 X = OR, NHR

O OH

O

O OH OH

HO O

HO

OH

HO X OH

H2N

O

*

*

(3R,4R ) : L-CBG-II (3S,4R ) : L-CBG-III (3R,4S ) : L-CBG-IV

O

Figure 10.1 Glu analogues prepared by AspAT-catalysed transamination

1 M.p. 178 C; ½25 D ¼ þ24:0 (c ¼ 1.3, 6 M HCl). H NMR (400 MHz, D2O) 3.83 (1H, dd, J ¼ 4.5 and 8.5 Hz), 2.58 (1H, m, J ¼ 5.0, 7.0, 8.5 Hz), 2.23 (1H, ddd, J ¼ 5.0, 8.5 and 14.0 Hz), 1.96 (1H, ddd, J ¼ 5.0, 8.5, 13.5 Hz), 1.25 (3H, d, J ¼ 7.0 Hz); 13C NMR (100 MHz, D2O) 185.3, 178.5, 53.9, 39.5, 36.9, 17.8. Anal. (C6H11NO4) C, H, N: calc., 44.72, 6.88, 8.69; found, 44.60, 6.94, 8.64.

10.3.2

Conclusion

AspAT has been shown to display a broad substrate spectrum. This chemoenzymatic procedure is, therefore, a very convenient way to prepare a variety of L-2,4-syn Glu analogues substituted at the 4-position by alkyl7 or functionalized substituents.12 Moreover, this catalyst has been used for the preparation of 4,4-disubstituted10 and (2S,3R)-3-methyl9 Glu derivatives, as well as the cyclobutane analogues LCBG II–IV.8 The different Glu analogues prepared to date using this methodology are reported in Figure 10.1.

References and Notes 1. Hwang, B.-Y., Cho, B.-K., Yun, H., Koteshwar, K. and Kim, B.-G., Revisit of aminotransferase in the genomic era and its application to biocatalysis. J. Mol. Catal. B: Enzym., 2005, 37, 47–55. 2. Ager, D.J., Li, T., Pantaleone, D.P., Senkpeil, R.F., Taylor, P.P. and Fotheringham, I.G., Novel biosynthetic routes to non-proteinogenic amino acids as chiral pharmaceutical intermediates. J. Mol. Catal. B: Enzym., 2001, 11, 199–205. 3. Yun, H., Cho, B.-K. and Kim, B.-G., Kinetic resolution of (R,S)-sec-butylamine using omegatransaminase from Vibrio fluvialis JS17 under reduced pressure. Biotechnol. Bioeng., 2004, 87, 772–778. 4. Shin, J.S. and Kim, B.G., Comparison of the o-transaminases from different microorganisms and application to production of chiral amines. Biosci. Biotechnol. Biochem., 2001, 65, 1782–1788.

10.3 Unnatural Amino Acids by Enzymatic Transamination

309

5. Iwasaki, A., Yamada, Y., Ikenaka, Y. and Hasegawa, J., Microbial synthesis of (R)- and (S)-3,4dimethoxyamphetamines through stereoselective transamination. Biotechnol. Lett., 2003, 25, 1843–1846. 6. Yun, H., Lim, S., Cho, B.-K. and Kim, B.-G., o-Amino acid:pyruvate transaminase from Alcaligenes denitrificans Y2k-2: a new catalyst for kinetic resolution of -amino acids and amines. Appl. Environ. Microbiol., 2004, 70, 2529–2534. 7. 1 was prepared in three steps from commercially available methyl 3-hydroxy-2-methylenebutyrate and triethyl orthoacetate (Aldrich): Alaux, S., Kusk, M., Sagot, E., Bolte, J., Jensen, A.A., Brauner-Osborne, H., Gefflaut, T. and Bunch, L., Chemoenzymatic synthesis of a series of 4substituted glutamate analogues and pharmacological characterization at human glutamate transporters subtypes 13. J. Med. Chem., 2005, 48, 7980–7992. 8. Faure, S., Jensen, A.A., Maurat, V., Gu, X., Sagot, E., Aitken, D.J., Bolte, J., Gefflaut, T. and Bunch, L., Stereoselective chemoenzymatic synthesis of the four stereoisomers of L-2-(2carboxycyclobutyl)glycine and pharmacological characterization at human excitatory amino acid transporter subtypes 1, 2, and 3. J. Med. Chem., 2006, 49, 6532–6538. 9. Xian, M., Alaux, S., Sagot, E. and Gefflaut, T., Chemoenzymatic synthesis of glutamic acid analogues: substrate specificity and synthetic applications of branched chain aminotransferase from Escherichia coli. J. Org. Chem., 2007, 72, 7560–7566. 10. Helaine, V., Rossi, J., Gefflaut, T., Alaux, S. and Bolte, J., Synthesis of 4,4-disubstituted Lglutamic acids by enzymatic transamination. Adv. Synth. Catal., 2001, 343, 692–697. 11. Acetaldehyde is used to limit enzyme inhibition by trapping SO2 produced from CSA. It can be omitted if Escherichia coli AspAT is used instead of pig heart enzyme, the bacterial enzyme being less sensitive to inhibition by SO2. 12. (a) Sagot, E., Jensen, A.A., Pickering, D., Stensbol, T.B., Nielsen, B., Assaf, Z., Aboab, B., Bolte, J., Gefflaut, T. and Bunch L., Chemo-enzymatic synthesis of (2S,4R)-2-amino-4-(3-(2,2diphenylethylamino)-3-oxopropyl)pentanedioic acid: a novel selective inhibitor of human excitatory amino acid transporter subtype 2. J. Med. Chem., 2008, 51, 4085–4092. (b) Sagot, E., Jensen, A.A., Pickering, D., Stensbol, T.B., Nielsen, B., Chapelet, M., Bolte, J., Gefflaut, T. and Bunch L., Chemo-enzymatic synthesis of a series of 2,4-syn-functionalized (S)glutamate analogues: new insight into the structure–activity relation of ionotropic glutamate receptor subtypes 5, 6, and 7. J. Med. Chem., 2008, 51, 4093–4103.

310


10.4

Synthesis of L-Pipecolic Acid with D1-Piperidine-2-carboxylate Reductase from Pseudomonas putida Hisaaki Mihara and Nobuyoshi Esaki

L-Pipecolic acid, a key component of many antibiotic and anticancer biomolecules,

1

serves as an important chiral pharmaceutical intermediate. We have developed an enzymecoupled system consisting of D1-piperidine-2-carboxylate reductase (Pip2C) from Pseudomonas putida, glucose dehydrogenase (GDH) from Bacillus subtilis, and L-lysine -oxidase from Trichoderma viride, affording L-pipecolic acid from L-lysine in high yield with an excellent enantioselectivity (Figure 10.2).2 10.4.1 10.4.1.1 • • • • • • • • • • • • • • • • •

Procedure 1: Preparation of Enzymatic Crude Extract Materials and Equipment

Bacto-tryptone (1 g) bacto-yeast extract (0.5 g) NaCl (1 g) NaOH (5 M) deionized water ampicillin (100 mg mL1, sterilized through a 0.22 mm filter) chloramphenicol (25 mg mL1 in ethanol) isopropyl--D-thiogalactopyranoside (1 M, sterilized through a 0.22 mm filter) tris-HCl buffer (20 mM, pH 7.0) membrane disc filters with 0.22 mm pore size autoclave one 500 mL Sakaguchi flask with a poromeric silicone plug one 20 mL test tube with a poromeric silicone plug clean bench incubation shaker sonicator centrifuge.

NH2 L-Lysine α-oxidase

N

COOH

NADPH

Pip2C reductase H2N COOH

N H

COOH

Gluconolactone

GDH

NADP+

Glucose

90%, >99.7% ee

Figure 10.2 Enzyme-coupled system for synthesis of L-pipecolic acid from L-lysine

10.4 Synthesis of L-Pipecolic Acid with D1-Piperidine-2-carboxylate Reductase

10.4.1.2

311

Procedure

1. Bacto-tryptone (1 g), bacto-yeast extract (0.5 g) and NaCl (1 g) were dissolved in 95 mL of deionized water and the pH was adjusted to 7.0 with 5 M NaOH. The volume was adjusted to 100 mL withdeionized water.A portion ofthe resulting solution(5 mL)wasplacedin a 20 mL test tube with a poromeric silicone plug and the rest was placed in a 500 mL Sakaguchi flask with a poromeric silicone plug. The media were sterilized by autoclaving (121 C, 20 min). 2. The 5 mL medium in a test tube was charged with ampicillin at 100 mg mL1 and chloramphenicol at 25 mg mL1 and inoculated with a single colony of recombinant Escherichia coli BL21(DE3) cells harbouring both pDPKA,3 which carries a gene for Pip2C reductase, and pSTVbsGDH,3 which has a B. subtilis GDH4 gene between the EcoRI and PstI sites of pSTV28. 3. The cells were shaken at 245 rpm for 20 h at 37 C. The preculture (100 mL) was transferred to the 100 mL medium containing 100 mg mL1 ampicillin and 25 mg mL1 chloramphenicol in a Sakaguchi flask and shaken at 245 rpm for 16 h at 37 C. 4. The culture was charged with 1 mM isopropyl--D-thiogalactopyranoside to induce gene expression and cultivated another 3 h. 5. The cells were harvested by centrifugation at 5000 rpm for 10 min at 4 C and washed twice with 20 mM tris-HCl (pH 7.0). The washed cells were disrupted by sonication for 1 min on ice. The supernatant was collected by centrifugation at 7500 rpm for 30 min at 4 C to obtain the cell-free crude extract. 10.4.2

Procedure 2: Synthesis of L-Pipecolic Acid

N H

10.4.2.1

COOH


• L-Lysine monohydrochloride (502 mg, 2.75 mmol) • glucose (990 mg, 5.5 mmol) • -nicotinamide adenine dinucleotide phosphate (NADPþ) sodium salt (4.87 mg, 2 nmol) • flavin adenine dinucleotide (FAD) disodium salt hydrate (8.66 mg, 10 nmol) • 100 mM tris-HCl buffer pH 7.5 • L-lysine -oxidase from T. viride (Seikagaku Corporation, 30 units) • catalase from bovine liver (Sigma–Aldrich, 500 units) • NaOH (10 M) • reciprocal shaker • one 100 mL flask with a poromeric silicone plug. 10.4.2.2

Procedure

1. To a 100 mL flask containing L-lysine (102 mg, 0.6 mmol), glucose (990 mg, 5.5 mmol), NADPþ (4.87 mg, 2.0 mmol), FAD (8.66 mg, 10.0 mmol), L-lysine -oxidase

312


(30 units), catalase from bovine liver (500 units) in 100 mM tris-HCl buffer solution pH 7.5 (10 mL) was added a crude extract of the recombinant E. coli BL21(DE3) cells with pDPKA and pSTVbsGDH (30 mg protein – as prepared above). The flask was stoppered with a poromeric silicone plug and shaken at 30 C for 26 h. 2. L-Lysine (100 mg portions) was added to the reaction mixture at 3, 6, 11, and 17 h intervals. To prevent a decrease in pH, the reaction mixture was adjusted to pH 7.5 with 10 M NaOH throughout the reaction course. 3. After 17 h, a titre of 210 mM (27 g L1) L-pipecolic acid was achieved with satisfactorily high optical purity (>99.7 % ee). The molar yield of L-pipecolic acid relative to L-lysine was 90 %. 4. L-Pipecolic acid can be isolated from the resultant reaction solution by commonly used methods, such as ion-exchange chromatography and crystallization, as described previously.5 Enantiomeric excess was determined by high-performance liquid chromatography with a Chiralpak WE column (4.6 mm 250 mm, Daicel Chemical Industries, Tokyo), 2 mM CuSO4, 0.75 mL min1, 50 C, monitored at 254 nm; L-pipecolic acid rt ¼ 14.7 min, D-pipecolic acid rt ¼ 18.0 min. 10.4.3

Conclusion

The procedure can provide a higher amount of L-pipecolic acid in a shorter reaction time than the previously reported system,6 indicating that it is applicable in industrial production of L-pipecolic acid. A similar system was successfully employed in the enzymatic synthesis of several cyclic amino acids by our group.7

References 1. (a) Germann, U.A., Shlyakhter, D., Mason, V.S., Zelle, R.E., Duffy, J.P., Galullo, V., Armistead, D.M., Saunders, J.O., Boger, J. and Harding, M.W., Cellular and biochemical characterization of VX-710 as a chemosensitizer: reversal of P-glycoprotein-mediated multidrug resistance in vitro. Anti-Cancer Drugs, 1997, 8, 125. (b) Vezina, C., Kudelski, A. and Sehgal, S.N., Taxonomy of the producing streptomycete and isolation of the active principle. J. Antibiot. (Tokyo), 1975, 28, 721. (c) Lehmann, J., Hutchison, A.J., McPherson, S.E., Mondadori, C., Schmutz, M., Sinton, C.M., Tsai, C., Murphy, D.E., Steel, D.J., Williams, M., Cheney, D.L. and Wood, P.L., CGS 19755, a selective and competitive N-methyl-D-aspartate-type excitatory amino acid receptor antagonist. J. Pharmacol. Exp. Ther., 1988, 246, 65. (d) Boger, D.L., Chen, J.H. and Saionz, K.W., ()Sandramycin: total synthesis and characterization of DNA binding properties. J. Am. Chem. Soc., 1996, 118, 1629. (e) Hirota, A., Suzuki, A., Aizawa, K. and Tamura, S., Structure of Cyl-2, a novel cyclotetrapeptide from Cylindrocladium scoparium. Agric. Biol. Chem., 1973, 37, 955. 2. Muramatsu, H., Mihara, H., Yasuda, M., Ueda, M., Kurihara, T. and Esaki, N., Enzymatic synthesis of L-pipecolic acid by D1-piperidine-2-carboxylate reductase from Pseudomonas putida. Biosci. Biotechnol. Biochem., 2006, 70, 2296. 3. (a) Muramatsu, H., Mihara, H., Kakutani, R., Yasuda, M., Ueda, M., Kurihara, T. and Esaki, N., The putative malate/lactate dehydrogenase from Pseudomonas putida is an NADPH-dependent D1-piperidine-2-carboxylate/D1-pyrroline-2-carboxylate reductase involved in the catabolism of D-lysine and D-proline. J. Biol. Chem., 2005, 280, 5329. (b) Mihara, H., Muramatsu, H., Kakutani, R., Yasuda, M., Ueda, M., Kurihara, T. and Esaki, N., N-Methyl-L-amino acid dehydrogenase from Pseudomonas putida. FEBS J., 2005, 272, 1117.

10.4 Synthesis of L-Pipecolic Acid with D1-Piperidine-2-carboxylate Reductase

313

4. Lampel, K.A., Uratani, B., Chaudhry, G.R., Ramaley, R.F. and Rudikoff, S., Characterization of the developmentally regulated Bacillus subtilis glucose dehydrogenase gene. J. Bacteriol., 1986, 166, 238. 5. Rodwell, V.W., Pipecolic acid. Methods Enzymol. Pt 2, 1971, 17, 174. 6. (a) Fujii, T., Mukaihara, M., Agematu, H. and Tsunekawa, H., Biotransformation of L-lysine to Lpipecolic acid catalyzed by L-lysine 6-aminotransferase and pyrroline-5-carboxylate reductase. Biosci. Biotechnol. Biochem., 2002, 66, 622. (b) Fujii, T., Aritoku, Y., Agematu, H. and Tsunekawa, H., Increase in the rate of L-pipecolic acid production using lat-expressing Escherichia coli by lysP and yeiE amplification. Biosci. Biotechnol. Biochem. 2002, 66, 1981. 7. Yasuda, M., Ueda, M., Muramatsu, H., Mihara, H. and Esaki, N., Enzymatic synthesis of cyclic amino acids by N-methyl-L-amino acid dehydrogenase from Pseudomonas putida. Tetrahedron Asymmetry. 2006, 17, 1775.

314

10.5


Synthesis of Substituted Derivatives of L-Phenylalanine and of other Non-natural L-Amino Acids Using Engineered Mutants of Phenylalanine Dehydrogenase Philip Conway, Francesca Paradisi and Paul Engel

We have used a series of biocatalysts produced by site-directed mutations at the active site of L-phenylalanine dehydrogenase (PheDH) of Bacillus sphaericus,1,2 which expand the substrate specificity range beyond that of the wild-type enzyme, to catalyse oxidoreductions involving various non-natural L-amino acids. These may be produced by enantioselective enzyme-catalysed reductive amination of the corresponding 2-oxoacid.3,4 Since the reaction is reversible, these biocatalysts may also be used to effect a kinetic resolution of a 5 D,L racemic mixture. Here, we describe, as a representative example, an efficient chemical synthesis of 4fluorophenylpyruvic acid (Procedure 1, Section 10.5.1) followed by its biocatalytic conversion to L-4-fluorophenylalanine catalysed by the N145V mutant2–4 of PheDH (Procedure 2, Section 10.5.2). 10.5.1

Procedure 1: Preparation of 4-Fluorophenylpyruvic Acid O

O +

HN

F

NH

O

O N H 130 °C

HN

F O

NH

OH

NaOH 5M 100°C

F

O 87% crude yield (8.4 g)

10.5.1.1 • • • • • • • • • • • • • • • • • •

O


Hydantoin (5.2 g, 52 mmol) 4-fluorobenzaldehyde (5 mL, 47 mmol) piperidine (9.9 mL, 100 mmol) cyclohexane ethyl acetate distilled water (200 mL) nitrogen gas HCl 12 M (20 mL) NaOH 5 M aqueous (21 mL) thin-layer chromatography (TLC) plates (silica gel 60 F254, Merck) Whatman pH indicator paper type CF (1–14 range) two-necked reaction flask equipped with a magnetic stirrer bar, 100 mL magnetic stirrer and heating plate equipment for reflux condenser oil bath filter paper glass pipettes one 250 mL separatory funnel

yield 73%

10.5 Synthesis of Substituted Derivatives of L-Phenylalanine

315

• flask • rotary evaporator. 10.5.1.2

Procedure

1. Hydantoin (5.2 g, 52 mmol) was added to piperidine (9.9 mL, 100 mmol) in a twonecked reaction flask equipped with a magnetic stirrer bar and heated to 130 C under nitrogen flux. 4-Fluorobenzaldehyde (5 mL, 47 mmol) was added dropwise to the stirring mixture. The reaction was monitored by TLC (eluent: ethyl acetate/cyclohexane, 1:4) and reached completion in 30 min. Attention. At room temperature hydantoin is insoluble in piperidine, but it will dissolve at approximately 80 C. Nitrogen is required to remove any traces of oxygen, but the reaction does not need to be moisture-free. 2. The reaction mixture was cooled to 60 C and water (200 mL) was added and stirred vigorously for 30 min. A yellow, tarry side product precipitated and was removed by filtration on filter paper. 3. The filtrate was acidified with HCl 12 M (approximately 20 mL) to pH 2.0, monitored with Whatman pH indicator paper. The resulting precipitate was collected on filter paper and dried under vacuum to afford 4-fluorobenzyl hydantoin (8.4 g, 87 %). 1 H NMR (500 MHz; DMSO) 6.41 (s, 1H, olefin), 11.43–10.32 (m, 2H, NHs), 7.22 (t, J ¼ 8.76 Hz, 2H, Aromatic), 7.66 (dd, J ¼ 7.38, 5.99 Hz, 2H, Aromatic). 4. A fraction of the crude 4-fluorobenzyl-hydantoin (1 g, 4.9 mmol) was mixed with a solution of NaOH 5 M (21 mL) in a two-necked reaction flask equipped with a magnetic stirrer bar. The mixture was refluxed at 100 C for 2.5 h. The reaction showed a strong colour change as it progressed. As the starting material was added to the base, the mixture turned a bright orange colour, which then lightened as the final product was formed. 5. The reaction was allowed to cool to room temperature and HCl 12 M was added dropwise to generate the free acid. The 4-fluorophenylpyruvic acid was extracted with EtOAc and the organic layer was dried with MgSO4 (which was then removed by passing through filter paper) and evaporated in vacuo. 4-Fluorophenylpyruvic acid was obtained as a yellow solid (0.7 g, 73 %). 1 H NMR (500 MHz; CDCl3) 1.37 (3H, t, J ¼ 7.1 Hz, CH3), 4.35 (2H, q, J ¼ 7.1 Hz, CH2), 6.48 (1H, s, H-3), 6.64 (1H, bs, OH), 7.01–7.07 (2H, m, Ar-H), 7.71–7.78 (2H, m, Ar-H).

10.5.2

Procedure 2: Enzymatic Synthesis of L-4-Fluorophenylalanine F

F O

H 2O

NH3 ONa O

NH 2 OH

(S)

PheDH mutant NAD +

NADH YADH CH3 CHO

EtOH

O yield 0.19 g (86%) >99% ee

316


10.5.2.1


• • • • • • • • • • • • • • • • •

4-Fluorophenylpyruvate sodium salt (204 mg, 1 mmol) nicotinamide adenine dinucleotide (NADþ, 12 mg, 20 mmol) KCl (76 mg, 1 mmol) HCOONH4 (252 mg, 4 mmol) ethylenediaminetetraacetic acid (EDTA, 4 mg, 10 mmol) EtOH (0.5 mL) tris buffer 50 mM, pH 8.5 (10 mL) yeast alcohol dehydrogenase (ADH, 1 mg, 633 U mg1) PheDH N145V mutant HCl 6 M (2 mL) NH4OH aqueous as needed HCl 1 M (150 mL) ninhydrin 15 mL plastic tubes with sealable cap orbital shaker incubator centrifuge high-performance liquid chromatograph fitted with chiral column such as CHIROBIOTIC T • Dowex monosphere 550A (OH) anion-exchange resin (60 mL) • equipment for ion-exchange chromatography • rotary evaporator. 10.5.2.2

Procedure

1. 4-Fluorophenylpyruvate sodium salt (204 mg, 1 mmol) was mixed with NADþ (12 mg, 20 mmol), KCl (76 mg, 1 mmol), HCOONH4 (252 mg, 4 mmol) and EDTA (4 mg, 10 mmol). EtOH (0.5 mL) and tris buffer 50 mM pH 8.5 (10 mL) were used to dissolve the reagents. Attention. Despite the use of the oxo acid in the salt form, and the addition of EtOH, the non-natural oxoacid substrates are not fully soluble at 0.1 M. Greater dilution resulted in lower conversion rates, however. 2. Reaction was initiated with 1 mg yeast ADH (663 U mg1) plus 10 mg N145V and the mixture was held at 25 C in an orbital shaker incubator. 3. Amino acid formation was monitored over 24 h by chiral high-performance liquid chromatography (CHIROBIOTIC T column) with samples diluted 10-fold (in H2O) to a suitable concentration. 4. The reaction was quenched by adding HCl 6 M (2 mL). After centrifugation, the crude reaction mixture (precipitate and supernatant) was ready for purification. 5. The crude reaction mixture was brought to pH 12 with aqueous NH3 and applied to a 60 mL column of Dowex monosphere 550A (OH) anion exchanger. Inorganic salts were eluted with 300 mL water. Subsequently, the amino acid was eluted with 1 M HCl (150 mL) and detected with ninhydrin. The eluate was concentrated in vacuo to give the pure amino acid as a white solid (0.19 g, 86 %).

10.5 Synthesis of Substituted Derivatives of L-Phenylalanine

10.5.3

317

Conclusion

The hydantoin route for synthesis of the 2-oxoacid6 has been performed with a variety of starting aldehydes and appears to work more reliably than the method described earlier.3 In spite of the restricted solubility of the intermediate oxoacid substrate for the biocatalytic step, the reaction proceeds to a good final overall yield with more substrate dissolving to replace that consumed in the reaction. The combined procedure appears to be quite versatile.

References 1. Seah, S.Y.K., Britton, K.L., Rice, D.W., Asano, Y. and Engel, P.C., Single amino acid substitution in Bacillus sphaericus phenylalanine dehydrogenase dramatically increases its discrimination between phenylalanine and tyrosine substrates. Biochemistry, 2002, 41, 11390. 2. Seah, S.Y.K., Britton, K.L., Rice, D.W., Asano, Y. and Engel, P.C., Kinetic analysis of phenylalanine dehydrogenase mutants designed for aliphatic amino acid dehydrogenase activity with guidance from homology-based modelling. Eur. J. Biochem., 2003, 270, 4628. 3. Busca, P., Paradisi, F., Moynihan, E., Maguire, A.R. and Engel, P.C., Enantioselective synthesis of non-natural amino acids using phenylalanine dehydrogenases modified by site-directed mutagenesis. Org. Biomol. Chem., 2004, 2, 2684. 4. Paradisi, F., Collins, S., Maguire, A.R. and Engel, P.C., Phenylalanine dehydrogenase mutants: efficient biocatalysts for synthesis of non-natural phenylalanine derivatives. J. Biotechnol., 2007, 128, 408. 5. Paradisi, F., Conway, P.A., Maguire, A.R. and Engel, P.C., Engineered dehydrogenase biocatalysts for non-natural amino acids: efficient isolation of the D-enantiomer from racemic mixtures. Org. Biomol. Chem., 2008, 6, 3611. 6. Billek, G., p-Hydroxyphenylpyruvic acid. Org. Synth. Collect. Vol., 1973, 5, 627.

11 Enzymatic Oxidation Chemistry 11.1

Monoamine Oxidase-catalysed Reactions: Application Towards the Chemo-enzymatic Deracemization of the Alkaloid (–)-Crispine A Andrew J. Ellis, Renate Reiss, Timothy J. Snape and Nicholas J. Turner

Previously, we reported a general method for the chemo-enzymatic deracemization of primary,1 secondary2 and tertiary3 amines in high yield and enantiomeric excess. The deracemization process involves a two-step, one-pot reaction and employs an enantioselective amine oxidase (MAO-N) in combination with a nonselective chemical reducing agent (Figure 11.1). We have further demonstrated the utility of this variant enzyme by way of the asymmetric synthesis of the natural product (þ)-crispine A in 97 % ee.4 The previously reported MAON-D5 variant, which contains five important mutations (Ile246Met/Asn336Ser/Met348Lys/ Thr384Asn/Asp385Ser) was used; its preparation has been described previously.5,6

R1

N

R3

R2 R4

enantio selective amine oxidase R1

(S) NH3:BH3 1

R

2

N

R

R3

N+

R2 R4

4

R3

R (R)

Figure 11.1 Enzymatic deracemization of racemic amines via a two-step, one-pot process utilizing an enantioselective amine oxidase in combination with ammonia–borane.



320

Enzymatic Oxidation Chemistry

11.1.1 11.1.1.1 • • • • • • • • • • •

Procedure 1: Preparation of the Biocatalyst Materials and Equipment

aqueous suspension of Escherichia coli BL21 competent cells (50 mL – Invitrogen) plasmid pET16b (Novagen) containing the variant MAO-N-D5 gene (1 mL)5,6 Luria–Bertani (LB) broth containing 100 mg mL1 ampicillin LB agar containing 100 mg mL1 ampicillin (contained in Petri dishes) potassium phosphate buffer (K2HPO4–KH2PO4) (50 mM, pH 7.6) Erlenmeyer flask (250 mL) with foam bung ice-bath centrifuge shaker/incubator static incubator Falcon tube (50 mL).

11.1.1.2

Procedure

1. The plasmid was transformed into E. coli BL21 competent cells as per the manufacturer’s instructions (Invitrogen). 2. The transformed cell suspension (50 mL) was spread onto an LB-ampicillin agar plate and incubated at 37 C for 16 h. 3. A single colony was used to inoculate 5 mL of LB-ampicillin broth contained in a 50 mL Falcon tube. This was incubated at 37 C, 250 rpm for 3 h. 4. The cell suspension (130 mL) was used to inoculate 25 mL of fresh LB-ampicillin broth contained in a 250 mL Erlenmeyer flask with a foam bung. This culture was incubated at 30 C, 250 rpm for 24 h. 5. Cells were harvested by centrifugation at 3000 Gav for 30 min. 6. Cell pellets were washed by resuspension in potassium phosphate buffer (10 mL) and were subjected to further centrifugation at 3000 Gav. 7. Wet cell pellets were then used directly in Procedure 2 (Section 11.1.2). 11.1.2

Procedure 2: Deracemization of (–)-Crispine A MeO

MAO-N-D5 N

MeO H

BH3 .NH3 buffer (pH7.6)

MeO N

MeO H

43%, 97% ee

11.1.2.1


• whole wet cells expressing the MAO-N-D5 variant protein (440 mg) • potassium phosphate buffer (K2HPO4–KH2PO4) (2.46 mL, 0.1 M, pH 7.6) • racemic crispine A (6.0 mg, 0.03 mmol)

11.1 Monoamine Oxidase-catalysed Reactions

• • • • • • • •

321

ammonia–borane complex (3.5 mg, 0.11 mmol) dichloromethane anhydrous MgSO4 glass vial (10 mL) sealed with a stopper shaker/incubator (set to 30 C, 250 rpm) microcentrifuge 0.2 mm in-line syringe filter rotary evaporator.

11.1.2.2

Procedure

8. Whole wet cells (440 mg) expressing the MAO-N-D5 variant enzyme were suspended in 0.1 M potassium phosphate buffer pH 7.6 (2.46 mL). Racemic crispine A (6.0 mg, 0.03 mmol) was added to this suspension followed by ammonia–borane (3.5 mg, 0.11 mmol). The vial was sealed with the stopper and the mixture was incubated at 30 C, 250 rpm and samples (0.5 mL) taken periodically for analysis. 9. For high-performance liquid chromatography (HPLC) analysis: samples (0.5 mL) were clarified by centrifugation at 14 000 Gav for 5 min and the supernatant was decanted, filtered through a 0.2 mm in-line syringe filter and analysed directly by chiral HPLC (see below). 10. For isolated material: the reaction mixture was clarified by centrifugation at 14 000 Gav for 5 min and the supernatant decanted and extracted with dichloromethane. The dichloromethane phase was dried (MgSO4) and concentrated in vacuo to yield the title compound as a colourless oil (4 mg, 43 %). 1 H NMR (CDCl3): d 1.67–1.76 (1H, m), 1.83–1.95 (2H, m), 2.29–2.37 (1H, m), 2.64 (1H, q, J 8.5), 2.66–2.71 (1H, m), 2.73 (1H, br. dt, J 16.1, 3.8), 2.93–3.00 (1H, m), 3.02–3.07 (1H, m), 3.12–3.17 (1H, m), 3.51 (1H, br. t, J 6.0), 3.82 (6H, s), 6.54 (1H, s), 6.58 (1H, s). 13 C NMR (CDCl3): d 22.1, 27.6, 30.5, 48.1, 53.0 (CH2), 55.8, 55.9 (CH3), 62.6, 108.7, 111.1 (CH), 125.8, 130.2, 147.2, 147.3 (C). Electrospray ionization mass spectrometry (þve): found m/z 234.1 (MHþ, 100%). []D ¼ þ88.4 (c ¼ 1.0, CHCl3). Enantiomeric excess was determined by HPLC with an OD-H column (90 % isohexane in isopropanol), 1.0 mL min1, 210 nm; major enantiomer Rt ¼ 18.6 min, 97 % ee. 11.1.3

Conclusion

The procedure is very easy to perform and is highly reproducible and may be applied to a wide range of substrates; see below for selected examples:

NH

95%, 99% ee

H N

80%, 98% ee

N

75%, 99% ee

322


References 1. Alexeeva, M., Enright, A., Dawson, M.J., Mahmoudian, M. and Turner, N.J., Deracemization of -methylbenzylamine using an enzyme obtained by in vitro evolution. Angew. Chem. Int. Ed., 2002, 41, 3177. 2. Carr, R., Alexeeva, M., Dawson, M.J., Gotor-Fernandez, V., Humphrey, C.E., Turner, N.J., Directed evolution of an amine oxidase for the preparative deracemisation of cyclic secondary amines. ChemBioChem, 2005, 6, 637. 3. Dunsmore, C.J., Carr, R., Fleming, T. and Turner, N.J., A chemo-enzymatic route to enantiomerically pure cyclic tertiary amines. J. Am. Chem. Soc., 2006, 128, 2224. 4. Bailey, K.R., Ellis, A.J., Reiss, R., Snape, T.J. and Turner, N.J., A template-based mnemonic for monoamine oxidase (MAO-N) catalyzed reactions and its application to the chemo-enzymatic deracemisation of the alkaloid (–)-crispine A. Chem. Commun., 2007, 3640. 5. Alexeeva, M., Carr, R. and Turner, N.J., Directed evolution of enzymes: new biocatalysts for asymmetric synthesis. Org. Biomol. Chem., 2003, 1, 4133. 6. Carr, R., Alexeeva, M., Enright, A., Eve, T.S.C., Dawson, M.J. and Turner, N.J., Directed evolution of an amine oxidase possessing both broad substrate specificity and high enantioselectivity. Angew. Chem. Int. Ed., 2003, 42, 4807.

11.2 Glucose Oxidase-catalysed Synthesis of Aldonic Acids

11.2

323

Glucose Oxidase-catalysed Synthesis of Aldonic Acids Fabio Pezzotti, Helene Therisod and Michel Therisod

The classical chemical synthesis of aldonic acids makes use of stoichiometric amounts of bromine, copper or silver hydroxides, or mercuric acetate in totally nonecologically acceptable processes.1,2 Glucose oxidase (EC 1.1.3.4) has the reputation of being extremely specific for D-glucose (a characteristic commonly used in analytical biochemistry). However, by extending the reaction time and the amount of enzyme, we were able to prepare gram-scale quantities of xylonic, galactonic, mannonic, 2-deoxygluconic and 2-aminogluconic acids.3,4 This was made possible by the availability of glucose oxidase, produced on an industrial scale and at low cost by Novozymes. The reaction takes place in water under atmospheric oxygen (as an oxidant). The products are isolated in pure form either by precipitation (2-aminogluconic acid) or by filtration through an ion-exchange resin. 11.2.1

Procedure: Synthesis of Xylonic, Galactonic, Mannonic, 2-Deoxygluconic Acid and Synthesis of 2-Amino-2-deoxy-gluconic Acid (Glucosaminic Acid) Glucose oxidase

O

H 2O/NaOH (pH-stat)

O HO

HO OH O2, H 2O

H 2O2

O

Dowex 1 (AcO–) elution HCl

OH HO

Aldonolactone

Aldose

COOH Aldonic acid

Catalase

Glucose oxidase

O

HO

OH

OH

OH HO

H 2 O/NaOH

O

(pH-stat)

HO

HO N H2

OH

O 2 , H2O

H2 O 2

Glucosamine Catalase

11.2.1.1

N H2

O

HO HO

OH COO – N H3+ Glucosaminic acid (precipitates)


• Aldose (D-xylose, D-galactose, D-mannose, D-2-deoxyglucose, D-glucosamine hydrochloride) (11.1 mmol) • glucose oxidase (Gluczyme, from Novozymes) (200 mg, 400 U) • catalase* (Catazyme, from Novozymes) (1 mL, 25 kU) • 1 M sodium hydroxide (11.1 mmol) • 1 M hydrochloric acid • Dowex 1 (acetate form) • pH-stat • rotary evaporator. *The use of catalase is not essential, as long as large amounts of glucose oxidase are used, since this last enzyme apparently is quite resistant to high concentration of hydrogen

324


peroxide. Moreover, Gluczyme contains some catalase activity. Catalase may be more useful if another source of purified glucose oxidase is used. 11.2.1.2

Procedure

11. The aldose (11.1 mmol) was dissolved in water to a final concentration of 0.5 M and subjected to oxidation by addition of glucose oxidase (200 mg, 400 U) and a large excess of catalase (1 mL, 25 kU). The mixture was vigorously stirred under air and the pH was kept constant at 7.5 by means of a pH-stat adding continuously 1 M NaOH. The conversion degree was directly calculated considering the volume of added 1 M NaOH, since 1 mol of NaOH neutralizes 1 mol of aldonic acid formed. 12. The reaction mixture was then filtered through a Dowex 1 (AcO) column to eliminate the enzymes and any residual substrate. 13. The aldonic acid (as a mixture of lactones) was then recovered by elution with 1 M aqueous HCl and evaporation in vacuo. 14. For the synthesis of 2-aminogluconic acid, the starting glucosamine hydrochloride was first neutralized by 1 M NaOH before starting the enzymatic oxidation. The insoluble product was recovered by concentration of the reaction medium and filtration. 15. The identity and purity of each product were confirmed by 1H and 13C NMR spectroscopy (D2O–Na2CO3). 11.2.1.3

2-Deoxy-D-gluconic Acid

Yield 92 %. 1 H NMR (360 MHz, D2O): d 2.52 (dd, 1H, J 15, J 5.8, H2), 2.55 (dd, 1H, J 15, J 8.3, H20 ), 3.51 (dd, 1H, J 8.3, J 2, H4), 3.76 (dd,1H, J 12, J 6.3, H6), 3.8 (m, 1H, H5), 3.9 (dd, 1H, J 11.5, J 2.9, H60 ), 4.3 (ddd, 1H, J 8.3, J 5.4, J 1.8, H3). 13 C NMR (62 MHz, D2O): d 41.61 C2, 63.07 C6, 67.89 C3, 71.28 C5, 72.87 C4, 180.33 C1 (in accordance with literature data5). []D ¼ þ5.05 (c ¼ 2.18, H2O) (lit. 6 []D ¼ þ4.2). 11.2.1.4

D-Galactonic

Acid

Yield 77 %. 1 H NMR (360 MHz, D2O): d 3.56 (dd, 1H, J 9.6, J 1.5, H4), 3.62 (d, 2H, J 6.4, H6–H60 ), 3.88 (dd, 1H, J 9.5, J 1.5, H5), 3.90 (dd, 1H, J 9.5, J 1.5, H3), 4.2 (d, 1H, J 1.5, H2). 13 C NMR (62 MHz, D2O): d 63.35 C6, 69.82 C4, 70.16 C5, 71.42 C3, 71.57 C2, 179.64 C1 (in accordance with literature data7). []D ¼ þ1.6 (c ¼ 10, H2O) (lit.8 []D ¼ þ0.4). 11.2.1.5

D-Mannonic

Acid

Yield 70 %. 1 H NMR (360 MHz, D2O): d 3.6 (dd, 1H, J 11.5, J 2.7, H6), 3.67 (bs, 2H, H5–H4), 3.75 (d, 1H, J 11.5, H60 ), 3.92 (d, 1H, J 5.6, H3), 4.06 (d, 1H, J 5.6, H2). 13 C NMR (62 MHz, D2O): d 62.95 C6, 70.42 C3, 70.49 C5, 70.86 C4, 73.84 C2, 179.25 C1 (in accordance with literature data9). []D ¼ 8.9 (c ¼ 10, H2O) (lit.8 []D ¼ 8.8).

11.2 Glucose Oxidase-catalysed Synthesis of Aldonic Acids

11.2.1.6

D-Xylonic

325

Acid

Yield 90 %. 1 H NMR (360 MHz, D2O): d 3.50 (dd, 1H, J 11.8, J 5.4, H5), 3.62 (dd, 1H, J 11.7, J 3.9, H50 ), 3.71 (m, 1H, H4), 3.76 (dd, 1H, J 5.9, J 2.5, H3), 3.99 (d, 1H, J 2.55, H2), 3.84 (d, 1H, J 2.4). 13 C NMR (62 MHz, D2O): d 62.72 C5, 72.54 C3, 73.09 C2, 73.19 C4, 179.19 C1 (in accordance with literature data10). []D ¼ þ7.05 (c ¼ 10, H2O) (lit. 11 []D ¼ þ7.4). 11.2.1.7

D-Glucosaminic

Acid

Yield 76 %. 1 H NMR (360 MHz, D2O): d 3.05 (d, 1H, J 5.4, H2), 3.28 (dd, 1H, J 10.4, J 4.7, H6), 3.3–3.4 (m, 2H, H4, H5), 3.42 (dd, 1H, J 10.4, J 4.3, H60 ), 3.57 (dd, 1H, J 5.4, J 1.8, H3). 13 C NMR (62 MHz, D2O): d 59.54 C2 , 62.92 C6, 72.18 C3, 72.97 C5, 73.8 C4, 181.11 C1 (in accordance with literature data12). []D ¼ 15 (c ¼ 4, 2.5 % aqueous HCl) (lit.13 []D ¼ 14) 11.2.2

Conclusion

We have devised a very simple procedure for the preparative synthesis of various aldonic acids from the corresponding aldoses. This ‘green chemistry’ process takes advantage of the availability of cheap, robust industrial enzymes.

References 1. (a) Varela, O., Oxidative reactions and degradations of sugars and polysaccharides. Adv. Carbohydr. Chem. Biochem., 2003, 58, 307. (b) DeLederkremer, R.M. and Marino, C., Acids and other products of oxidation of sugars. Adv. Carbohydr. Chem. Biochem., 2003, 58, 199. 2. Pringsheim, H. and Ruschman, G., Preparation of glucosaminic acid. Ber. Dtsch. Chem. Ges. 1915, 48, 680. 3. Pezzotti, F., Therisod, H. and Therisod, M., Enzymatic synthesis of D-glucosaminic acid from D-glucosamine. Carbohydr. Res., 2005, 340, 139. 4. Pezzotti, F. and Therisod, M., Enzymatic synthesis of aldonic acids. Carbohydr. Res., 2006, 341, 2290. 5. Freimund, S., Huwig, A., Giffhorn, F. and Ko¨pper, S., Rare keto-aldoses from enzymatic oxidation: substrates and oxidation products of pyranose 2-oxidase. Chem. Eur. J., 1998, 4, 2442. 6. Horton, D. and Philips, K.D., Diazo derivatives of sugars. Synthesis of methyl 2-deoxy-2-diazoD-arabino-hexonate, its behaviour on photolysis and thermolysis, and conversion into a pyrazole derivative. Carbohydr. Res., 1972, 22, 151. 7. Ramos, M.L., Caldeira, M.M. and Gil, V., NMR study of the complexation of D-galactonic acid with tungsten (VI) and molybdenum (VI). Carbohydr. Res., 1997, 297, 191. 8. Levene, P.A., The specific rotations of hexonic and 2-amino-hexonic acids and of their sodium salts. J. Biol. Chem., 1924, 59, 123. 9. Horton, D., Walaszek, Z. and Ekiel, I., Conformations of D-gluconic, D-mannonic, and Dgalactonic acids in solution, as determined by n.m.r. spectroscopy. Carbohydr. Res., 1983, 119, 263. 10. Serianni, A.S., Nunez, H.A. and Barker, R., Cyanohydrin synthesis: studies with carbon-13labeled cyanide. J. Org. Chem., 1980, 45, 3329.

326


¨ ber die Bestandtheile des Mais-Marks und des Hollander11. Browne, C.A. and Tollens, B., U Marks und das gleichzeitige Vorkommen von Araban und Xylan in den Pflanzen. Berl. Dtsch. Chem. Ges., 1902, 35, 1457. 12. Horton, D., Thomson, J.K., Varela, O., Nin, A. and Lederkremer, R.M., Confirmation of the structures of the products obtained on acylation of 2-amino-2-deoxy-D-gluconic acid. Carbohydr. Res., 1989, 193, 49. 13. Hope, D.B. and Kent, P.W., Ester and lactone linkages in acidic polysaccharides. Part II. Lactones of D-glucosaminic acid. J. Chem. Soc. Abstr., 1955, 1831.

11.3 Oxidation and Halo-hydroxylation of Monoterpenes

11.3

327

Oxidation and Halo-hydroxylation of Monoterpenes with Chloroperoxidase from Leptoxyphium fumago Bjoern-Arne Kaup, Umberto Piantini, Matthias Wu¨st and Jens Schrader

Chloroperoxidase (CPO) from Leptoxyphium fumago (formerly Caldariomyces fumago) is a unique enzyme showing broad substrate specificity and featuring industrially relevant reactions, including halogenation and oxidation reactions. Recently, monoterpenes were discovered to be substrates for both oxidation (Figure 11.2) and halo-hydroxylation (Figure 11.3) by CPO in the absence and presence of halide ions respectively.1 In the case of halo-hydroxylation of (1S)-(þ)-3-carene, excellent ee values of >99 % were detected. The introduction of two stereogenic centres in one step makes this reaction very interesting given the fact that 3-carene has been investigated as starting material for the synthesis of different valuable target compounds, such as fragrances and -lactam antibiotics.2,3 11.3.1 11.3.1.1 • • • •

Procedure 1: Halo-hydroxylation of (1S)-(þ)-3-Carene by CPO Materials and Equipment

(1S)-(þ)-3-Carene (>99 %, 10 mM) sodium chloride, bromide or iodide (p.a., 10 mM) citric acid buffer (100 mM, pH 3.5) tert-butanol (>99 %)

CH 2 OH

CHO CPO +H2 O 2

Geraniol

Geranial

Figure 11.2 Oxidation of the monoterpene alcohol geraniol to geranial by CPO in the presence of hydrogen peroxide and absence of halide ions.

HO X

CPO +H 2O2 + X

(1S)-(+)-3-Carene

–

(1S,3R,4R,6R )-4-Halo3,7,7-trimethyl-bicyclo[4.1.0]heptane-3-ol

Figure 11.3 Stereoselective halo-hydroxylation of the monoterpene hydrocarbon (1S)-(þ)-3carene by CPO in the presence of hydrogen peroxide and halide ions (X ¼ Cl, Br or I).

328

• • • • • • •


CPO (0.045 mg mL1, 1.2 mM) Hydrogen peroxide (1 M) n-hexane (>99 %) sodium sulfate (p.a.) one 50 mL vessel with screw cap magnetic stir bar magnetic stirrer.

11.3.1.2

Procedure

16. For the conversion of (1S)-(þ)-3-carene approximately 0.045 mg mL1 (1.2 mM) CPO was incubated in 100 mM citric acid buffer, pH 3.5 with 25 % (v/v) tert-butanol containing 10 mM (1S)-(þ)-3-carene (final assay concentration) and 10 mM sodium chloride, sodium bromide or sodium iodide (final assay concentrations) in a 50 mL vessel on a magnetic stirrer (300 rpm) at room temperature. Hydrogen peroxide was added to a total concentration of 10 mM over a reaction time of 60 min at a rate of 165 mM min1 (165 mM portions every minute). 17. Samples were extracted with n-hexane, dried over sodium sulfate and stored at 20 C until analysed by coupled gas chromatography–mass spectrometry (GC-MS). GC–MS: Shimadzu GC-17A/GCMS-OP5050 system; column: Valco Bond VB-S (30 m 0.25 mm 0.25 mm); injection: split 50:1 at 230 C, 1 mL; carrier gas: helium, 1.3 mL min1; interface temperature: 250 C; oven program: starting temperature 50 C, rate 5 C min1 for 30 min to 200 C. Product identification was carried out by NMR analysis using 1H, 13C and 1H/1H correlation spectroscopy techniques.1 Under the given conditions, molar conversion yields were 90 % after 10 min, 90 % after 20 min and 60 % after 30 min for iodo-, bromoand chloro-halohydrin formation respectively. 11.3.2 11.3.2.1 • • • • • • • • • •

Procedure 2: Oxidation of Geraniol by CPO Materials and Equipment

Geraniol (96 %, 2 mM) citric acid buffer (100 mM, pH 3.5) tert-butanol (>99 %) CPO (0.2 mg mL1, 5 mM) hydrogen peroxide (1 M) n-hexane (>99 %) sodium sulfate (p.a.) one 50 mL vessel with screw cap magnetic stir bar magnetic stirrer.

11.3.2.2

Procedure

18. For conversion of geraniol approximately 0.2 mg mL1 (5 mM) CPO was incubated in 100 mM citric acid buffer, pH 3.5, with 25 % (v/v) tert-butanol containing 2 mM

11.3 Oxidation and Halo-hydroxylation of Monoterpenes

329

geraniol (final assay concentration) in a 50 mL vessel on a magnetic stirrer (300 rpm) at room temperature. Hydrogen peroxide was added to a total concentration of 2 mM over a reaction time of 60 min at a rate of 33 mM min1 (100 mM portions every 3 min). 19. Samples were extracted with n-hexane, dried over sodium sulfate and stored at 20 C until GC–MS analysis. GC–MS: Shimadzu GC-17A/GCMS-OP5050 system; column: Valco Bond VB-S (30 m 0.25 mm 0.25 mm); injection: split 50:1 at 230 C, 1 mL; carrier gas: helium, 1.3 mL min1; interface temperature: 250 C; oven program: starting temperature 50 C, rate 5 C min1 for 30 min to 200 C. The product was identified by comparison of its retention time and mass spectrum with those of a commercial standard substance. Under the given conditions, the final molar conversion yield was 37.5 %. 11.3.3

Conclusion

The procedures are very easy to reproduce and to scale up. Reaction products are isolated by evaporation of the extraction solvent (e.g. hexane, pentane). In the case of the carene halohydrin, further product purification is not necessary if reaction is allowed to proceed until total substrate conversion due to the high selectivity of product formation.

References 1. Kaup, B.A., Piantini, U., Wust, M. and Schrader, J., Monoterpenes as novel substrates for oxidation and halo-hydroxylation with chloroperoxidase from Caldariomyces fumago. Appl. Microbiol. Biotechnol., 2007, 73, 1087–1096. 2. Bhawal, B.M., Joshi, S.N., Krishnaswamy, D. and Deshmukh, A.R., (þ)-3-Carene, an efficient chiral pool for the diastereoselective synthesis of -lactams. J. Indian Inst. Sci., 2001, 81, 265–276. 3. Lochynski, S., Kowalska, K. and Wawrzenczyk, C., Synthesis and odour characteristics of new derivatives from the carane system. Flavour Fragr. J., 2002, 3, 181–186.

330

11.4


Chloroperoxidase-catalyzed Oxidation of Phenyl Methylsulfide in Ionic Liquids Cinzia Chiappe

The heme enzyme chloroperoxidase (CPO), produced by the marine fungus Caldariomyces fumago, is a versatile enzyme which exhibits a broad spectrum of chemical reactivities and it is recognized as a most promising biocatalyst for synthetic applications.1 Recently, pure (R)-phenyl methylsulfoxide (ee > 99 %) was prepared by chemo- and stereo-selective oxidation of phenyl methylsulfide with CPO in citrate buffer–ionic liquid mixtures.2 11.4.1

Procedure: Synthesis of (R)-Phenyl Methylsulfoxide O

O S

CH3

CPO

S

CH3

O S

+

CH3

H2O2 Ionic Liquid/ citrate buffer

11.4.1.1 • • • • • • • • • • • • •

95–100% e.e. >99%

5–0%


Thioanisole (6.2 mg, 50 mmol) CPO (67.4 U) hydrogen peroxide solution 7 wt% in water (50 mmol þ 25 mmol) sodium citrate buffer solution (0.1 M, pH 5) (1 mL) ionic liquid: cholinium citrate ([N1112OH][Citr], 1 mL) or 1,3-dimethylimidazolium methylsulfate ([mmim][MeSO4], 1 mL) or cholinium acetate ([N1112OH][OAc], 0.5 mL) sodium thiosulfate anhydrous magnesium sulfate diethyl ether filter paper one 10 mL test tube with a screw cap and equipped with a magnetic stirrer bar magnetic stirrer plate one 25 mL separatory funnel rotary evaporator.

11.4.1.2

Procedure

20. Thioanisole (6.2 mg, 50 mmol) and CPO (67.4 U) were magnetically stirred at room temperature in a 10 mL test tube for 5 min in 2 mL of a mixture of ionic liquid–sodium citrate buffer solution (0.1 M, pH 5). A 1:1 mixture was used in the case of [N1112OH][Citr] and [mmim][CH3SO4], whereas a 0.6:1.4 mixture was employed in the case of [N1112OH][OAc]. Hydrogen peroxide solution (7 wt%) was added in two portions: initially 50 mmol and then an additional 25 mmol after 2 h. After 4 h the reaction was quenched by the addition of excess Na2S2O3.

11.4 Chloroperoxidase-catalyzed Oxidation of Phenyl Methylsulfide

331

Table 11.1 Oxidation of thioanisole with hydrogen peroxide and CPO at room temperature in a 1:1 ionic liquid/citrate buffer Ionic liquid

Amount (v/v) of IL in citrate buffer (%)

Conversion (%)

Products Sulfoxide/ sulfone

[N1112OH][Citr] [mmim][CH3SO4] [N1112OH][OAc]

50 50 30

48 76 42

95:5 98:2 100:0

Ee (%) >99 (R) >99 (R) >99 (R)

21. The reaction mixture was extracted with diethyl ether. The organic portion was collected, dried over anhydrous magnesium sulfate and concentrated using a rotary evaporator. The crude product was analysed by gas chromatography on a chiral 30 m Chiraldex GTA column (helium flow 50 kPa, evaporator and detector set at 200 C, column temperature 90 C for 1 min, 8 C min1 to 170 C) after addition of anisole as an internal standard. The reaction performed in the 1:1 mixture of [mmim][CH3SO4]/sodium citrate buffer has been scaled up 50 times without any change in chemo- and enantio-selectivity. The crude reaction mixture was purified by chromatography on silica gel (hexane/ethyl acetate 6:1–1:1) to give pure phenyl methylsulfoxide (yield 70 %, ee 99 %). 11.4.2

Conclusions

Ionic liquids can be used as co-solvents for CPO-catalysed sulfoxidation. Table 11.1 gives details about different ionic liquids. The procedure is very easy to reproduce and the oxidation of thioanisole proceeds with high chemo- and stereo-selectivity.

References 1. Dembitsky, M.V., Oxidation, epoxidation and sulfoxidation reactions catalysed by haloperoxidases. Tetrahedron, 2003, 59, 4701. 2. Chiappe C., Neri, L. and Pieraccini, D., Application of hydrophilic ionic liquids as co-solvents in chloroperoxidase catalyzed oxidations. Tetrahedron Lett., 2006, 47, 5089.

332


Stereoselective Synthesis of b-Hydroxy Sulfoxides Catalyzed by Cyclohexanone Monooxygenase

11.5

Stefano Colonna, Nicoletta Gaggero, Sara Pellegrino and Francesca Zambianchi Chiral -hydroxy sulfoxides are well known for their usefulness as chiral auxiliaries for the preparation of a great variety of compounds, such as biaryl sulfoxides,1a cyclic sulfides,1b benzoxathiepines,1c benzothiazepines,1d allylic alcohols,1e macrolides1f and leucotrienes.1g The most straightforward method for their synthesis is the selective oxidation of the parent sulfides. Although a variety of catalytic systems2 have been introduced for this kind of oxidation, many of these methodologies are associated with several disadvantages, such as environmentally unfriendly catalysts, volatile organic solvents, harsh reaction conditions and low stereoselectivities. Therefore, the development of an environmentally benign, high yielding and clean approach for the synthesis of -hydroxy sulfoxides is needed. We have found that these goals can be achieved through the direct oxidation of different -hydroxy sulfides using cyclohexanone monooxygenase from Acinetobacter calcoaceticus (CHMO) as catalyst.3 CHMO is a bacterial Baeyer– Villigerase that has been used for a chemoenzymatic synthesis of a variety of key chiral products. This enzyme catalyzes the Baeyer–Villiger oxidation of cyclohexanone with formation of the corresponding E-caprolactone; the only reagents consumed are O2 and reduced nicotinamide adenine dinucleotide phosphate (NADPH). Apart from cyclohexanone and other ketones and aldehydes, CHMO can oxidize a wide series of organic compounds containing electron-rich heteroatoms, i.e. sulfides are converted to sulfoxides,4 sulfites to sulfates,5 selenides to selenoxides,6 tertiary amines to N-oxides7 and phospines to phospinoxides.8 In many cases these reactions are highly enantioselective9 (Figure 11.4).

S

S

1R,2R,RS n

O

OH

n

O

1S,2S,SS

1-3a major diasteroisomer

OH

S CHMO/G6PDH n

OH

G6P/NADP/O2

(±)-trans -1-3 1R,2R,SS

S n

S O

OH

n

O

OH

1S,2S,RS 1-3b minor diasteroisomer

Figure 11.4 Oxidation of -hydroxy sulfides to -hydroxy sulfoxides catalyzed by CHMO

11.5 Stereoselective Synthesis of -Hydroxy Sulfoxides

11.5.1 11.5.1.1 • • • • • • • • •

333

Procedure 1: Preparation and Purification of CHMO Materials and Equipment

Buffer A: 0.02 M potassium phosphate buffer with 0.01 M dithiothreitol, pH 7.2 (NH4)2SO4 NaCl NADPþ sonicator centrifuge Fractogel EMD DEAE, 10 cm 2 cm ion-exchange column Matrex gel red A, agarose 5 %; 10 cm 2 cm affinity column lyophilizor.

11.5.1.2

Procedure

1. A. calcoaceticus was grown as described by Trudgill10 and the transformed microorganism was cultivated essentially as described previously by Doig et al.11 2. All steps of purification were carried out at 4 C using buffer A. 3. The cells obtained from 1 L of culture medium were harvested, disrupted by sonication and cell debris removed by centrifugation. 4. The supernatant was subjected to fractionation with (NH4)2SO4 and the fraction which precipitated between 40 and 85 % saturation was recovered by centrifugation at 6000g for 30 min. 5. The pellet was redissolved in buffer A, dialysed overnight against the same buffer and loaded on an anion-exchange column (Fractogel EMD DEAE, 10 cm 2 cm) which was previously equilibrated with buffer A. The enzyme was eluted with a linear gradient from 0 to 0.15 M NaCl for 30 min in the same buffer, at a flow rate of 2 mL min1. 6. Active fractions were collected and loaded on an affinity column (Matrex gel red A, agarose 5 %; 10 cm 2 cm) previously equilibrated with buffer A and eluted with the same buffer but containing NADPþ 0.05 M (flow rate of 1 mL min1). Active fractions were collected, dialysed overnight against buffer A and lyophilized. 11.5.2 11.5.2.1

Procedure 2: Oxidation of b-Hydroxy Sulfides to b-Hydroxy Sulfoxides Catalyzed by CHMO Materials and Equipment

• Sulfides (–)-1–3 (1 mg) • CHMO (1 U) is not that commercially available12 (see Procedure 1, Section 11.5.1) • glucose-6-phosphate dehydrogenase from Leuconostoc mesenteroides (G6PDH) (18 U mL1) • glucose-6-phosphate (G6P) (50 mM, 15 mg) • NADPþ (0.5 mM, 1 mg) • pH 8.6 tris-HCl buffer solution 50 mM (1 mL) • diethyl ether (1 mL) • propan-2-ol (1 mL)

334

• • • • • • • • • •


n-hexane ethyl acetate silica gel 60 (230-400 mesh) reaction flask equipped with a magnetic stirrer bar magnetic stirrer plate separatory funnel rotary evaporator high-performance liquid chromatography (HPLC) equipment Chiracel OD column (Daicel, Illkirch, France) column chromatography equipment.

11.5.2.2

Procedure

1. Sulfide (–)-1 or (–)-2 or (–)-3 (1 mg) was added to 50 mM tris-HCl buffer pH 8.6 (1 mL) containing NADPþ (1 mg), G6P (15 mg), partially purified CHMO (1 U) and G6PDH (18 U). The reaction mixture was gently stirred at 25 C (see Table 11.2 for reaction times). Table 11.2

Oxidation of racemic -hydroxy sulfides catalyzed by CHMO

Sulfide Time (h) C (%)a

ee sulfidesa (%) [E]

eea (%)

dr major/ minora Major

(–)-1b,c n¼0 (–)-2d,e n¼1 (–)-2d,e n¼1 (–)-3f,g n¼2 (–)-3f,g n¼2 a

Minor

24

97

69 [1.6]

83:17

53

91

1

36

47 [261]

>99:1

98

-

5

47

87 [299]

99:1

98 (1S,2S,SS)

95 (1R,2R,SS)

1

52

50 [4.3]

89:11

63

56

3

78

79 [3.3]

82:18

45

78

Determined by HPLC analysis on Chiralcel OD column. trans-2-(Phenylsulfinyl)cyclopentan-1-ol (1a). Major diastereoisomer. M.p. 102 C. IR (KBr): 3390, 3058, 1651, 1085, 1028 cm1. 1H NMR (CDCl3, 200 MHz): d 1.62–1.83 (m, 5 H, 30 -H, 4-H, 5-H), 2.07 (m, 1 H, 300 -H), 3.02 (m, 2 H, 2-H, OH), 4.64 (m, 1 H, 1-H), 7.50 (m, 3 H, Ar-H), 7.77 (m, 2 H, Ar-H) ppm. Electron-ionization mass spectrometry (EI-MS): m/z (%) ¼ 210 (100) [M]þ. c trans-2-(Phenylsulfinyl)cyclopentan-1-ol (1b). Minor diastereoisomer. M.p. 97 C. IR (KBr): 3295, 3058, 1637, 1085, 1012 cm1. 1H NMR (CDCl3, 200 MHz): d 1.53–1.71 (m, 4 H, 4-H, 5-H), 1.84–2.06 (m, 3 H, 3-H, OH), 3.03 (m, 1 H, 2-H), 4.56 (m, 1 H, 1-H), 7.85 (m, 5 H, Ar-H) ppm. EI-MS: m/z (%) ¼ 210 (100) [M]þ. d trans-2-(Phenylsulfinyl)cyclohexan-1-ol (2a). Major diastereoisomer (1S,2S,SS). M.p. 156–157 C. IR (KBr): 3444, 2931, 1628, 1077, 1002 cm–1. 1H NMR (CDCl3, 300 MHz): d 1.11–1.40 (m, 6 H, 4-H, 5-H, 6-H), 1.73 (m, 2 H, 3-H), 2.75 (m, 1 H, 2-H), 3.03 (m, 1 H, 1-H), 4.16 (br s, 1 H, OH), 7.55 (m, 3 H, Ar-H), 7.74 (m, 2 H, Ar-H) ppm. EI-MS: m/z (%) ¼ 224 (100) [M]þ. ½20 D ¼þ134 (c ¼ 1.5, CHCl3). e trans-2-(Phenylsulfinyl)cyclohexan-1-ol (2b). Minor diastereoisomer (1R,2R,SS). M.p. 138–139 C. IR (KBr): 3450, 2931, 1648, 1077, 1012 cm1. 1H NMR (CDCl3, 300 MHz): d 1.09–1.71 (m, 7 H, 30 -H, 4-H, 5-H, 6-H), 2.11 (m, 1 H, 300 H), 2.66 (m, 1 H, 2-H), 3.93 (m, 1 H, 1-H), 4.01 (br s, 1 H, OH), 7.58 (m, 5 H, Ar-H) ppm. EI-MS: m/z (%) ¼ 224 (100) [M]þ. ½20 D ¼þ100:9 (c ¼ 1.5, CHCl3). f trans-2-(Phenylsulfinyl)cycloheptan-1-ol (3a). Major diastereoisomer. M.p. 148 C. IR (KBr): 3353, 3059, 1579, 1050, 1014 cm1. 1H NMR (CDCl3, 200 MHz): d 1.19–1.90 (m, 10 H, 3-H, 4-H, 5-H, 6-H, 7-H), 2.85 (br s, 1 H, OH), 2.94 (m, 1 H, 2-H), 4.32 (m, 1 H, 1-H), 7.56 (m, 3 H, Ar-H), 7.76 (m, 2 H,Ar-H) ppm. EI-MS: m/z (%) ¼ 238 (100) [M]þ. g trans-2-(Phenylsulfinyl)cycloheptan-1-ol (3b). Minor diastereoisomer. M.p. 125 C. IR (KBr): 3358, 3057, 1638, 1050, 1014 cm1. 1H NMR (CDCl3, 200 MHz): d 1.24–1.89 (m, 10 H, 3-H, 4-H, 5-H, 6-H, 7-H), 2.16 (br s, 1 H, OH), 2.99 (m, 1 H, 2-H), 4.16 (m, 1 H, 1-H), 7.53 (m, 3 H, Ar-H), 7.88 (m, 2 H, Ar-H) ppm. EI-MS: m/z (%) ¼ 238 (100) [M]þ. b

11.5 Stereoselective Synthesis of -Hydroxy Sulfoxides

335

2. The reaction medium was extracted with an equal volume of diethyl ether (1 mL), evaporated, diluted to original volume with propan-2-ol (1 mL) and analysed by chiral HPLC (Chiralcel OD column: rate flow 1 mL min1, l 254 nm; for (–)-1: n-hexane/ propan-2-ol, 90/10; for (–)-2: n-hexane/propan-2-ol, 95/5; for (–)-3: n-hexane/propan2-ol, 90/10) in order to evaluate the degree of oxidation and the enantiomeric excess. 3. The crude reaction mixture was purified by column chromatography (n-hexane/ethyl acetate: for (–)-1, 2/8; for (–)-2, 4/6; for (–)-3, 4/6) affording pure diastereomers 1a–3a and 1b–3b. Enantiomeric excess (ee), conversion C and diastereomeric ratio (dr) are reported in Table 11.2. 11.5.3

Conclusion

The kinetic resolution of -hydroxy sulfides mediated by CHMO provides an excellent result in the case of sulfide (–)-2 and moderate results with (–)-1 and (–)-3. Indeed, the enzyme-catalysed oxidation to sulfoxide 2a showed remarkable enantio- and diastereoselectivity with an enantiomeric ratio E of 299 and with an ee 98 % (C ¼ 47 %).

References 1. (a) Broutin, P.E. and Colobert, F., Enantiopure -hydroxysulfoxide derivatives as novel chiral auxiliaries in asymmetric biaryl Suzuki reactions. Org. Lett., 2003, 5, 3281. (b) Eames, J. and Warren, S., Synthesis of cyclic sulfides and allylic sulfides by phenylsulfanyl (PhS-) migration of -hydroxy sulfides. J. Chem. Soc. Perkin Trans. 1, 1999, 2783. (c) Gelebe, A.C. and Kaye, P.T., Benzodiazepine analogues. Part 15. Synthesis of benzoxathiepine derivatives. Synth. Commun., 1996, 26, 4459. (d) Schwatz, A., Madan, P.B., Mohacsi, E., O’Brien, J.P., Todaro, L.J. and Coffen, D.L., Enantioselective synthesis of calcium channel blockers of the diltiazem group. J. Org. Chem., 1992, 57, 851. (e) Kesavan, V., Bonnet-Delpon, D. and Be´gue´, J.P., Fluoro alcohol as reaction medium: one-pot synthesis of -hydroxy sulfoxides from epoxides. Tetrahedron Lett., 2000, 41, 2895. (f) Solladie`, G., Almario, A. and Dominguez, C., Asymmetric synthesis of natural products monitored by chiral sulfoxides. Pure Appl. Chem., 1994, 66, 2159. (g) Corey, E.J., Clark, D.A., Goto, G., Marfat, A., Mioskowski, C., Samuelsson, B. and Hammarstrom, S., Stereospecific total synthesis of a ‘slow reacting substance’ of anaphylaxis, leukotriene C-1. J. Am. Chem. Soc., 1980, 102, 1436. 2. (a) Conte, V., Di Furia, F., Licini, G., Modena, G., Sbampato, G. and Valle, G., Enantioselective oxidation of -hydroxythioethers. Synthesis of optically active alcohols and epoxides. Tetrahedron Asymm., 1991, 2, 257. (b) Pitchen, P. and Kagan, H.B., An efficient asymmetric oxidation of sulfides to sulfoxides. Tetrahedron Lett., 1984, 25, 1049. 3. Colonna, S., Pironti, V., Zambianchi, F., Ottolina, G., Gaggero, N. and Celentano, G., Diastereoselective synthesis of -hydroxy sulfoxides: enzymatic and biomimetic approaches. Eur. J. Org. Chem., 2007, 363. 4. Colonna, S., Gaggero, N., Pasta, P., Ottolina G., Enantioselective oxidation of sulfides to sulfoxides catalysed by bacterial cyclohexanone monooxygenases. Chem. Commun., 1996, 2303. 5. Colonna, S., Gaggero, N., Carrea, G. and Pasta, P., Oxidation of organic cyclic sulfites to sulfates: a new reaction catalyzed by cyclohexanone monooxygenase. Chem. Commun., 1998, 415. 6. Branchaud, P. and Walsh C.T., Functional group diversity in enzymatic oxygenation reactions catalyzed by bacterial flavin-containing cyclohexanone oxygenase J. Am. Chem. Soc., 1995, 107, 2153, and references cited therein. 7. Ottolina, G., Bianchi, S., Belloni, B., Carrea, G. and Danieli, B., First asymmetric oxidation of tertiary amines by cyclohexanone monooxygenase. Tetrahedron Lett., 1999, 40, 8483.

336


8. Alphand, V., Archelas, A. and Furstoss, R., Microbial transformations 16. One-step synthesis of a pivotal prostaglandin chiral synthon via a highly enantioselective microbiological Baeyer– Villiger-type reaction. Tetrahedron Lett., 1989, 30, 3663. 9. Colonna, S., Pironti, V., Carrea, G., Pasta, P. and Zambianchi, F., Oxidation of secondary amines by molecular oxygen and cyclohexanone monooxygenase. Tetrahedron, 2004, 60, 569. 10. Trudgill, P.W., Cyclohexanone 1,2-monooxygenase from Acinetobacter NCIMB 9871. Methods Enzymol., 1990, 188, 70. 11. Doig, S.D., O’Sullivan, L.M., Patel, S., Ward, J.M. and Woodley, J.M., Large scale production of cyclohexanone monooxygenase from Escherichia coli TOP10 pQR239. Enzyme Microb. Technol., 2001, 28, 265. 12. Secundo, F., Zambianchi, F., Crippa, G., Carrea, G. and 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.

11.6 Enantioselective Kinetic Resolution of Racemic 3-Phenylbutan-2-one

11.6

337

Enantioselective Kinetic Resolution of Racemic 3-Phenylbutan-2one Using a Baeyer–Villiger Monooxygenase Anett Kirschner and Uwe T. Bornscheuer*

Baeyer–Villiger monooxygenases (BVMOs) mimic the chemical Baeyer–Villiger oxidation and belong to the class of oxidoreductases. Using molecular oxygen, they can convert ketones into esters or lactones.1 Most stereoselective Baeyer–Villiger oxidations were described for mono- and bi-cyclic ketones.2 Recently, we have shown that aliphatic acyclic3 and arylaliphatic4 ketones are also enantioselectively converted by a BVMO from Pseudomonas fluorescens DSM 50106, which was recombinantly expressed in Escherichia coli.5 Using whole cells of E. coli JM109 pGro7 pJOE4072.6 expressing this BVMO, preparative kinetic resolution of racemic 3-phenylbutan-2-one and subsequent hydrolysis of the ester product was performed giving (R)-3-phenylbutan-2-one in 45 % yield with 80 % ee and (S)-1-phenylethanol in 35 % yield and 93 % ee. 11.6.1 11.6.1.1 • • • • • • • • • • • • • • •

Procedure 1: Recombinant Expression of the BVMO from P. fluorescens DSM 50106 in E. coli Materials and Equipment

Tryptone (5 g) yeast extract (2.5 g) NaCl (5 g) distilled water ampicillin stock solution (100 mg mL1) chloramphenicol stock solution (50 mg mL1) L-rhamnose solution (20 % w/v) 1 L-arabinose solution (50 mg mL ) stored culture of E. coli JM109 harboring the chaperone plasmid pGro7 and the BVMOexpression plasmid pJOE4072.6 phosphate buffer solution (50 mM, pH 7.5) one 100 mL shake flask with a cotton plug one 1 L shake flask with a cotton plug shaker photometer centrifuge.

11.6.1.2

Procedure

1. Tryptone (5 g), yeast extract (2.5 g) and NaCl (5 g) were dissolved in distilled water, the volume was adjusted to 500 mL and then autoclaved (20 min, 120 C). A small portion of this Luria–Bertani (LB) medium (10 mL) was placed into a sterile 100 mL shake flask and ampicillin and chloramphenicol solutions were added (LBampþcm) to final concentrations of 100 mg mL1 and 20 mg mL1 respectively. The solution was inoculated with E. coli JM109 pGro7 pJOE4072.6 and shaken overnight at 37 C and 200 rpm. This overnight culture (2 mL) was used to inoculate 200 mL LBampþcm in a

338


1 L shake flask supplemented with 0.5 mg mL1 L-arabinose. The culture was incubated at 37 C and 200 rpm to an optical density (OD) at 600 nm of 0.6, where expression of the recombinant BVMO was induced by the addition of 0.2 % (w/v) L-rhamnose. Expression was performed for 4 h at 30 C and 200 rpm. 2. Cells were then harvested by centrifugation for 20 min at 4400g and 4 C. The medium was removed and the cell pellet was washed once with 50 mL phosphate buffer solution and centrifuged again. The cells can be stored in the fridge for a few days or used directly for biotransformation. 11.6.2

Procedure 2: Kinetic Resolution of Racemic 3-Phenylbutan-2-one O O

BVMO + O

O O2 NADPH +H+

11.6.2.1 • • • • • • • • • • • •

H2O NADP+


Phosphate buffer solution (50 mM, pH 7.5) racemic 3-phenylbutan-2-one (0.15 g, 1 mmol) -cyclodextrin (0.07 g, 0.5 mmol) glucose solution (1 M, 4 mL) ethyl acetate anhydrous sodium sulfate one 1 L shake flask with a cotton plug shaker photometer centrifuge one separatory funnel rotary evaporator.

11.6.2.2

Procedure

1. The cell pellet of E. coli JM109 pGro7 pJOE4072.6 was resuspended in phosphate buffer to a final OD at 600 nm of around 20. To 100 mL of this suspension in a 1 L shake flask racemic 3-phenylbutan-2-one (0.15 g, 1 mmol), -cyclodextrin (0.07 g, 0.5 mmol) and 1 M glucose solution (2 mL) were added. The reaction mixture was incubated at 30 C and 220 rpm. After 4 h, further 1 M glucose solution (2 mL) was added. 2. After 6 h, the mixture was centrifuged to remove cells from the solution. The pellet was washed and the reaction solution was extracted several times with ethyl acetate. The combined organic layers were dried over anhydrous sodium sulfate and concentrated using a rotary evaporator.

11.6 Enantioselective Kinetic Resolution of Racemic 3-Phenylbutan-2-one

339

3. The crude product was analyzed by chiral GC (Hydrodex--3P column)4 revealing 46 % conversion with 80 % and 94 % enantiomeric excess of substrate and product respectively, corresponding to an E-value of 82. 11.6.3

Procedure 3: Enzymatic Hydrolysis of (S)-1-Phenylethyl Acetate O OH +

O

O

11.6.3.1 • • • • • • • • • • • • •

CAL-A H 2O

+ O


Phosphate buffer solution (50 mM, pH 7.5), 50 mL hexane, 10 mL Candida antarctica lipase A (CAL-A, Chirazyme L-5, C2), 100 mg ethyl acetate anhydrous sodium sulfate silica gel thin-layer chromatography plates (silica gel 60 F254) reaction flask, 250 mL water bath magnetic stirrer separatory funnel rotary evaporator equipment for column chromatography.

11.6.3.2

Procedure

1. The crude product after kinetic resolution of racemic phenylbutan-2-one was dissolved in 10 mL hexane and transferred to a reaction flask containing 100 mg CAL-A in 50 mL phosphate buffer. The reaction mixture was stirred for 24 h at 30 C. 2. The reaction mixture was extracted several times with ethyl acetate. The combined organic layers were dried over anhydrous sodium sulfate and concentrated using a rotary evaporator after filtration. 3. Purification by silica-gel column chromatography (eluent: hexane:ethyl acetate, 5:1) gave 43 mg (S)-1-phenylethanol (35 % yield, 93 % ee) and 67 mg (R)-3-phenylbutan2-one (45 % yield, 80 % ee). 11.6.4

Conclusion

Using the procedure described herein, a racemic arylaliphatic ketone could be efficiently resolved using a BVMO. Similar arylaliphatic substrates were also shown to be enantioselectively converted on an analytical scale by a phenylacetone monooxygenase from Thermobifida fusca and a 4-hydroxyacetophenone monooxygenase from P. fluorescens ACB with good to high enantioselectivities.

340


References 1. (a) Walsh, C.T. and Chen, Y.C.J., Enzymic Baeyer–Villiger oxidations by flavin-dependent monooxygenases. Angew. Chem. Int. Ed. Engl., 1988, 27, 333. (b) Mihovilovic, M.D., Mu¨ller, B. and Stanetty, P., Monooxygenase-mediated Baeyer–Villiger oxidations. Eur. J. Org. Chem. 2002, 3711. (c) Mihovilovic, M.D., Enzyme mediated Baeyer–Villiger oxidations. Curr. Org. Chem., 2006, 10, 1265. (d) Kamerbeek, N.M., Janssen, D.B., van Berkel, W.J.H. and Fraaije, M.W., Baeyer–Villiger monooxygenases, an emerging family of flavin-dependent biocatalysts. Adv. Synth. Catal. 2003, 345, 667. 2. (a) Mihovilovic, M.D., Rudroff, F., Gro¨tzl, B., Kapitan, P., Snajdrova, R., Rydz, J. and Mach, R., Family clustering of Baeyer–Villiger monooxygenases based on protein sequence and stereopreference. Angew. Chem. Int. Ed., 2005, 44, 3609. (b) Taschner, M.J., Black, D.J. and Chen, Q.Z., The enzymatic Baeyer-Villiger oxidation : a study of 4-substituted cyclohexanones. Tetrahedron Asymm., 1993, 4, 1387. (c) Fraaije, M.W., Wu, J., Heuts, D.P.H.M., van Hellemond, E.W., Spelberg, J.H.L. and Janssen, D.B., Discovery of a thermostable Baeyer– Villiger monooxygenase by genome mining. Appl. Microbiol. Biotechnol., 2005, 66, 393. 3. Kirschner, A. and Bornscheuer, U.T., Kinetic resolution of 4-hydroxy-2-ketones catalyzed by a Baeyer–Villiger monooxygenase. Angew. Chem. Int. Ed., 2006, 45, 7004. 4. Geitner, K., Kirschner, A., Rehdorf, J., Schmidt, M., Mihovilovic, M.D. and Bornscheuer, U.T., Enantioselective kinetic resolution of 3-phenyl-2-ketones using Baeyer–Villiger monooxygenases. Tetrahedron Asymm., 2007, 18, 892. 5. Kirschner, A., Altenbuchner, J. and Bornscheuer, U.T., Cloning, expression, and characterization of a Baeyer–Villiger monooxygenase from Pseudomonas fluorescens DSM 50106 in E. coli. Appl. Microbiol. Biotechnol. 2007, 73, 1065. 6. Rodrı`guez, C., de Gonzalo, G., Fraaije, M.W. and Gotor, V., Enzymatic kinetic resolution of racemic ketones catalyzed by Baeyer–Villiger monooxygenases. Tetrahedron: Asymm., 2007, 18, 1338.

11.7 Desymmetrization of 1-Methylbicyclo(3.3.0)octane-2,8-dione

11.7

341

Desymmetrization of 1-Methylbicyclo[3.3.0]octane-2,8-dione by the Retro-claisenase 6-Oxo Camphor Hydrolase Gideon Grogan* and Cheryl Hill

A range of symmetrical bicyclic -diketones can be converted to 2,3-disubstituted cycloalkanones in high yield with high diastereomeric and enantiomeric excess using a cell-free preparation of a retro-Claisenase enzyme, or -diketone hydrolase, the gene for which has been heterologously expressed in Escherichia coli.1 11.7.1 11.7.1.1 • • • • • • • • • • • • •

Procedure 1: Preparation of the Crude Enzyme Materials and Equipment

Plasmid pGG3 E. coli BL21 (DE3) Luria–Bertani (LB) agar stock solution of kanamycin (1 mL, 30 mg mL1) stock solution of isopropylthio--galactopyranoside (IPTG, 2 mL, 1 M) phosphate buffer pH 7.0 (1 L of 50 mM) sterile plastic Petri dishes 30 mL sterile plastic culture Sterilin bottles (or 50 mL Falcon tubes) orbital shaker with controlled temperature (37 C) 2 L baffled Erlenmeyer flasks centrifuge with capacity to centrifuge several hundred millilitres ultrasonicator liquid nitrogen for snap freezing.

11.7.1.2

Procedure

1. Plasmid pGG3 (a Novagen pET-26b vector into which had been ligated the gene encoding 6-oxo camphor hydrolase (OCH))1 was transformed into E. coli BL21 (DE3) and the recombinant strain maintained on LB agar plates containing 30 mg mL1 kanamycin. 2. A single colony was used to inoculate a 5 mL starter culture in LB medium with 30 mg mL1 kanamycin, which was grown overnight at 37 C. 3. The turbid culture was then used to inoculate 500 mL of LB medium containing 30 mg mL1 kanamycin in a 2 L flask. The organism was grown at 37 C until an optical density A600 ¼ 0.5. 4. In order to induce expression of the OCH gene, 500 mL of a 1 M solution of IPTG was then added and the organism incubated at 37 C for 3 h. 5. The culture was then centrifuged and the cell pellet resuspended in 50 mL 50 mM phosphate buffer pH 7. 6. The cell suspension was disrupted by ultrasonication and the cell debris removed by centrifugation. 7. The supernatant was then snap-frozen in liquid nitrogen in aliquots for use directly as the biocatalyst, which had a specific activity of approximately 9 U mL1.

342


11.7.2

Procedure 2: Desymmetrization of 1-Methylbicyclo[3.3.0]octane-2, 8-dione2 O

R

O

6-axo camphor hydrolase phosphate buffer pH 7.0

R = Me, Et, allyl, propargyl

11.7.2.1

O R CO2H


• Phosphate buffer pH 7.0, 75 mL • crude OCH preparation (225 U – approximately 25 mL of the preparation described above) • diketone substrate3 (100 mg) • 2 M hydrochloric acid (few drops) • ethyl acetate (150 mL) • anhydrous MgSO4 for drying • thin-layer chromatography (TLC) plates (silica gel 60 F254, Merck) • 250 mL round-bottomed flask with a magnetic stirrer bar • magnetic stirrer plate • filter paper • 250 mL separatory funnel • rotary evaporator. 11.7.2.2

Procedure

1. Transfer 75 mL of the phosphate buffer into a 250 mL round-bottomed flask. To this add the enzyme solution (25 mL) and stir for 10 min at room temperature. 2. Make up a solution of 100 mg of the diketone substrates in ethanol (2 mL) and add this dropwise to the stirred buffer. Stir the reaction at room temperature overnight. 3. Analyse the reaction by TLC in a solvent system consisting of 1:1 ethyl acetate/hexane. The substrate has an Rf of approximately 0.55, and the keto acid product appears at or just above the baseline. If substrate is still present, then add a further 5 mL of the enzyme preparation (45 U approximately) and continue stirring for 2 h at room temperature. 4. When TLC shows that the reaction is complete, acidify the mixture to pH 3.0 using a few drops of 2 M HCl. The enzyme will precipitate. At this stage, the extraction of the product is facilitated if the precipitated protein is removed by centrifugation. 5. Extract the clear supernatant with ethyl acetate (3 50 mL) and dry the combined organic fractions with anhydrous magnesium sulfate. Filter off the drying agent and remove the solvent in vacuo to yield the crude keto-acid product. For purposes of analysis, the crude keto-acid was then treated with trimethylsilyl diazomethane converted to afford 3-(2-methyl-3-oxo-cyclopentyl)-propionic acid methyl ester, Rf 1:1 petrol/ethyl acetate (0.55).

11.7 Desymmetrization of 1-Methylbicyclo(3.3.0)octane-2,8-dione

343

1

H NMR (400 MHz; CDCl3) d 3.66 (3 H, s, OCH3), 2.48–231 (3 H, m), 2.20–2.02 (4 H, m), 1.72–1.58 (2 H, m), 1.37–1.32 (1 H, m) and 1.06 (3 H, d, J 7.0, CH3). 13C NMR (400 MHz; CDCl3) d 220.5 (C¼O), 173.9 (CO2Me), 51.8 (OCH3), 50.4 (CH), 44.2 (CH), 37.3 (CH2), 31.9 (CH2) and 29.6 (CH2), 26.9 (CH2) and 12.6 (CH3). m/z (chemical ionization; NH3) 202 [100 %, (M þ NH4)þ]. [Found: (M þ NH4)þ, 202.1439 C10H16O3 requires M þ NH4, 202.1443]. The diastereomeric excess (de) and enantiomeric excess (ee) were determined by first converting the methyl ester to the diastereomeric acetal by acid-catalysed reaction with (2R,3R)-2,3-butanediol. The acetals were then analysed on a capillary GC HP5 column (30 m 0.32 mm 0.25 mm): injector 250 C; 320 C; column 130 C isothermal. The de was calculated to be 82 % and the ee >95 %.2 Table 11.3 Desymmetrization of 1-alkylbicyclo[3.3.0]octane2,8-diones by OCH R Me Et Allyl Propargyl

11.7.3

De (%)

Ee (%)

82 81 86 78

>95 >95 >95 91

Conclusion

The desymmetrization of 1-alkylbicyclo[3.3.0]octane-2,8-diones can be achieved in a facile coenzyme-independent enzymatic reaction in buffer. Alkyl chains in the 1-position of up to at least five carbon atoms are tolerated.2 The yields of the crude keto-acids are essentially quantitative, and the enantiotopic discrimination by the enzyme is usually excellent.4 Work remains to be done on the optimization of this biocatalyst with respect to protein stability and reaction engineering, but it remains a unique and intriguing possibility for the generation of interesting intermediates bearing multiple chiral centres.

References and Notes ˚ crystal 1. Whittingham, J.L., Turkenburg, J.P., Verma, C.S., Walsh, M.A. and Grogan, G., The 2-A structure of 6-oxo camphor hydrolase: new structural diversity in the crotonase superfamily. J. Biol. Chem., 2003, 278, 1744. 2. Hill, C.L., Verma, C.S. and Grogan, G., Desymmetrisations of 1-alkylbicyclo[3.3.0]octane-2,8diones by enzymatic retro-Claisen reaction yield optically enriched 2,3-substituted cyclopentanones. Adv. Synth. Catal. 2007, 349, 916. 3. A synthetic method for the preparation of the diketone substrates has been presented: Hill, C.L., McGrath, M., Hunt, T. and Grogan, G., A generic and reproducible route to homo- and heteroannular bicyclic -diketones via Knochel-type 1,4-conjugate additions to ,-unsaturated cycloalkenones. Synlett, 2006, 309. 4. A proposed mechanism for the reaction, and the molecular basis for enantiotopic discrimination, based on X-ray crystallographic studies of the enzyme, has been reported: Leonard, P.M. and Grogan, G., Structure of 6-oxo camphor hydrolase H122A mutant bound to its natural product, (2S,4S)--campholinic acid: mutant structure suggests an atypical mode of transition state binding for a crotonase homolog. J. Biol. Chem., 2004, 279, 31312.

344

11.8


Synthesis of Optically Pure Chiral Lactones by Cyclopentadecanone Monooxygenase-catalyzed Baeyer–Villiger Oxidations Shaozhao Wang, Jianzhong Yang and Peter C.K. Lau*

Baeyer–Villiger monooxygenases (BVMOs), typified by cyclohexanone or cyclopentanone monooxygenases derived from Acinetobacter sp. NCIMB 9871 and Comamonas (formerly Pseudomonas) sp. NCIMB 9872 respectively, have been shown to be useful reagents for the preparation of optically active lactones with high enantiomeric excess (ee) and yield.1–4 Biooxidation using BVMOs is among the 12 recommended green chemistry research areas in the pharmaceutical industry, avoiding such hazardous reagents as organic peracids, chlorinated solvents or metals that are otherwise used in the chemical Baeyer–Villiger reactions.5 We recently introduced a new recombinant BVMO, called cyclopentadecanone monooxygenase (CPDMO) of Pseudomonas origin, that is capable of lactone formation from a broad spectrum of cyclic ketones ranging in size from substituted C6 to C15 ring compounds. In many cases, excellent enantioselectivity for the preparation of optically pure chiral lactones was demonstrated in whole-cell biotransformation experiments.6 The following section describes the biotransformations of 4-substituted cyclohexanones, 4-t-butyl cyclohexanone in particular, and several prochiral substrates to the corresponding lactones in good yield and excellent ee by whole-cell CPDMO desymmetrization, with a simple solvent (ethyl acetate) extraction for product recovery (Figure 11.5). CPDMO was also found to have an excellent enantioselectivity (E > 200) as well as 99 % (S)-selectivity toward 2-methyl-cyclohexanone for the production of 7-methyl-2-oxepanone, a potentially valuable chiral building block (Figure 11.6). In the latter case, scale-up synthesis in a 3 L fermenter was demonstrated. 11.8.1 11.8.1.1

Procedure 1: Propagation of Engineered Escherichia coli Strain BL21(DE3)[pCD201] Materials and Equipment

• Luria–Bertani (LB) medium (tryptone peptone 10 g L1, yeast extract, 5 g L1, NaCl 5 g L1) • LB-ampicillin (100 mg mL1) plates • LB-ampicillin media • isopropyl--D-thiogalacto-pyranoside (IPTG) • 30 % v/v sterile glycerol, 3 mL • 50 mL Erlenmeyer flask • 10 (2.5 mL) Eppendorf tubes • shaker. O

O E. coli / CPDMO

R

O

R

R = CH 3, CH2CH3, C(CH3)3

Figure 11.5 Lactone formation from 4-substituted cyclohexanone catalyzed by E. coli whole cells expressing CPDMO

11.8 Synthesis of Optically Pure Chiral Lactones by Cyclopentadecanone

11.8.1.2

345

Procedure

1. The E. coli strain BL21(DE3)[pCD201] expressing an IPTG-inducible CPDMO activity was streaked from a frozen stock on LB-ampicillin plates and incubated at 30 C until colonies were 1–2 mm in size. Refer to Sambrook et al.7 concerning media preparation, etc. 2. One colony was used to inoculate 10 mL of a LB-ampicillin medium in a 50 mL Erlenmeyer flask and incubated at 30 C, 200 rpm overnight. 3. Sterile glycerol (30 % v/v) was added and the mixture was divided into 1.0 mL aliquots and stored in a 80 C freezer. 4. The control carrier strain BL21(DE3)(pSD80) containing the plasmid pSD80 vector only was propagated using the same protocol except that no ampicillin was used in the plates or medium. 11.8.2

Procedure 2: Synthesis of (S)-5-t-Butyl-2-oxepanone O O

H3C H3C

11.8.2.1 • • • • • • • • • • • • •

CH3


LB-ampicillin medium (90 mL) 20% glucose solution (10 mL) 4-t-butyl cyclohexanone (50 mg, 0.32 mmol) 100 mM IPTG stock solution (100 mL) -cyclodextrin (0.25 g) ethyl acetate ( 100 mL) anhydrous sodium sulfate hexane ethyl acetate 500 mL baffled Erlenmeyer flask 500 mL separatory funnel rotary evaporator shaker.

11.8.2.2

Procedure

1. One tube of stock culture (1 mL) was thawed in a warm hand and used to inoculate an LB-ampicillin medium (90 mL) supplemented with 20 % glucose solution (10 mL) in a 500 mL baffled Erlenmeyer flask. 2. The culture was incubated at 30 C, 200 rpm until the optical density at 600 nm (OD600) was approximately 0.5–0.7 (around 1.5 h).

346


3. 100 mM IPTG stock solution was added (1 mL per milliliter of medium, final concentration 0.1 mM) followed by the substrate 4-t-butyl cyclohexanone (50 mg, 0.32 mmol) and -cyclodextrin (0.25 g). 4. The mixture solution was agitated at 30 C at 200 rpm for 26 h until the reaction was finished. 5. The culture solution was extracted with ethyl acetate (3 100 mL). Combined extracts were washed once with brine and dried with anhydrous Na2SO4. The solvent was removed on a rotary evaporator and the residue was purified by flash chromatography over silica gel to afford the title compound as white crystals (36 mg, 65 % isolated yield). Chiral-phase gas chromatography (GC) showed >99 % ee, []D ¼ 36 (c ¼ 1.7, CHCl3). Electron impact mass spectrometry (EI-MS) (m/e): 171 (1 %, Mþ þ 1), 155 (3 %), 114 (100 %), 86 (90 %). 1 H NMR (CDCl3, 500 MHz) d 4.35 (1H, dd, J1 ¼ 7.8 Hz , J2 ¼ 5.6 Hz), 4.16 (1H, dd, J1 ¼ 12.6 Hz, J2 ¼ 10.7 Hz), 2.72 (1H, dd, J1 ¼ 14.7 Hz, J2 ¼ 13.1 Hz), 2.58 (1H, t, J ¼ 11.7 Hz), 2.05 (2H, m), 1.50 (1H, m), 1.32 (2H, m), 0.89 (9H, s) ppm. 13C NMR (CDCl3, 500 MHz) d 23.7, 27.3, 27.4, 27.5, 30.3, 32.9, 33.4, 50.7, 68.6, 176.2 ppm. 11.8.3

Procedure 3: Synthesis of Both Enantiomers of 7-Methyl-2-oxepanone

See Figure 11.6. 11.8.3.1 • • • • • • • •


Racemic 2-methyl cyclohexanone (100 mg, 0.89 mmol) LB-ampicillin medium (90 mL) 20% glucose solution (10 mL) 100 mM IPTG stock solution (10 mL) ethyl acetate (3 100 mL) hexane ethyl acetate anhydrous Na2SO4 (3 g) O O 36% O CH3

kinetic resolution with CPDMO reaction stopped at 50% conv, & separation

CH 3 99% ee O

O 19%

CH3

m -CPBA/TFA CH 2Cl 2

O CH3

99 % ee

99% ee

Figure 11.6 Kinetic resolution of 2-methycyclohexanone by CPDMO-catalyzed oxidation to yield both enantiomers of 7-methyl-2-oxepanone with high ee values.

11.8 Synthesis of Optically Pure Chiral Lactones by Cyclopentadecanone

• • • • • • • • • •

347

m-chloroperoxybenzoic acid (50 mg) trifluoroacetic acid (TFA, 0.2 mL) silica gel 60, 200–425 mesh, Fisher Scientific (15 g) thin-layer chromatography plates (silica gel 60 F254, Merck) flask equipped with a magnetic stirrer bar, 50 mL magnetic stirrer plate chiral-phase GC, -Dex 225 column (Supelco Inc.) 50 mL and 500 mL separatory funnels rotary evaporator equipment for column chromatography.

11.8.3.2

Procedure

1. One tube of stock culture (1 mL) was thawed in a warm hand and was used to inoculate an LB-ampicillin medium (90 mL) plus 20 % glucose solution (10 mL) in a 500 mL baffled Erlenmeyer flask. The culture was incubated at 30 C, 200 rpm until OD600 was approximately 0.5–0.7 (around 1.5 h). 2. 100 mM IPTG stock solution (10 mL) was added followed by the substrate 2-methyl cyclohexanone (100 mg, 0.89 mmol). 3. The mixture was agitated at 200 rpm at 30 C (to monitor the reaction, aliquots were extracted with ethyl acetate and the organic layer analyzed by chiral-phase GC). 4. The kinetic resolution of racemic substrate with CPDMO reaction was stopped at 50 % conversion and immediately extracted with ethyl acetate. Combined extracts were washed once with brine and dried with anhydrous Na2SO4. 5. The mixture of optically pure lactone and ketone solution was evaporated by rotary evaporator to dryness. The residue mixture was separated by flash chromatography over silica gel (hexane:ethyl acetate 5:1), eluted first as colorless oil (S)-lactone (41 mg, 36 % yield, 99 % ee, []D ¼ 16, c ¼ 10, in CH2Cl2), followed by (R)-ketone as colorless oil (19 mg, 19 % yield, 99 % ee). 6. An analytical sample of (R)-ketone was chemically oxidized with m-chloroperoxybenzoic acid (TFA, CH2Cl2) to give (R)-lactone with 99 % ee on chiral-phase GC without losing any optical purity. 1

H NMR (250 MHz; CDCl3) d 1.36(d, J ¼ 6.5 Hz), 1.62(m, 4H), 1.93(m, 2H), 2.65(m, 2H), 4.28(m, 1H) ppm. 13 C NMR (63 MHz; CDCl3) d 22.5, 22.9, 28.2, 35.0, 36.2, 76.8, 175.6 ppm. EI-MS: 128 (1 %, Mþ), 113 (2 %), 84 (95 %), 55 (100 %). 11.8.4

Procedure 4: Scale-up Synthesis of (S)-7-Methyl-2-oxepanone in a 3 L Fermenter O O CH3

348


11.8.4.1


• LB medium (100 mL) • sugar solution (200 g L1) • supplemented M9 medium (1 L) containing: 4.0 g Na2HPO4, 2.0 g KH2PO4, 3.0 g (NH4)2SO4, 0.5 g NaCl, 1.0 g casamino acid, 0.12 g MgSO4, 58.0 mg CaCl22H2O, 50.0 mg thiamine, 50.0 mg ampicillin, 6.0 mg FeSO47H2O, 20 g glucose, and 4.5 mL US trace element solution as described.8 • racemic 2-methyl cyclohexanone (20 g) • phosphate buffer (0.05 M, pH 7.2) • KOH solution (2 M) • antifoam (Mazu DF 204, BASF) • 100 mM IPTG (1 mL) • dodecane (0.2 mL) • ethyl acetate • anhydrous sodium sulfate • micro-centrifuge tubes, 1.5 mL • cuvette, 1 mL • fermenter (3 L, Biobundles, Applikon Inc., US) • ReactIRTM 4000 spectrometer (Mettler Toledo, ASI Applied Systems, USA) (optional) • chiral-GC -Dex 225 column (Supelco Inc.) • high-performance liquid chromatograph (Hewlett Packard, Hp 1047A) • spectrophotometer (Hitachi Model U3210) • orbital incubator shaker (30 C, New Brunswick Scientific Innova43) • Erlenmeyer flasks, 500 mL • refrigerators (4C and 80 C) • refrigerated centrifuge • separation funnel, 3000 mL • rotary evaporator. 11.8.4.2

Procedure

1. Preculture. Frozen stock cell (1 mL, E. coli BL21(DE3)[pCD201] stored at 80 C) was thawed and precultured in 100 mL of LB medium in a 500 mL Erlenmeyer flask. The rotation speed of the incubator shaker was controlled at 250 rpm and the culture incubated overnight at 30 C. 2. Culture and bioconversion. The precultured cells were recovered by centrifugation at 4 C and the cell pellet was inoculated into a 3 L fermenter (stirred tank with two Rushton turbine impellers and four baffles) containing 1.0 L of supplemented M9 medium. 3. The cell culture was carried out under the following conditions: temperature 30 C; pH 7.0 controlled by the addition of 2 M KOH. The fermenter was aerated at 1 vvm via a submerged sparger and the agitation rate was controlled between 600 and 1000 rpm in order to maintain the dissolved oxygen concentration above 20 % air saturation. Foaming was controlled by addition of antifoam (Mazu DF 204, BASF). 4. The dissolved oxygen tension (DOT), feed rate and KOH consumption were monitored. When the cell density reached an OD600 ¼ 1, IPTG (1 mL, 100 mM) was added to induce the CPDMO expression.

11.8 Synthesis of Optically Pure Chiral Lactones by Cyclopentadecanone Table 11.4

349

Baeyer–Villiger oxidation by recombinant CPDMO using Procedure 2

Substrate

Yield (%)

4-Methylcyclohexanone 4-Ethylcyclohexanone cis-2,6-Dimethyl cyclohexanone

54 74 74

Enantioselectivity

Product

99% ee (S) 99% ee (S) 99% ee (?)

5-Methyl-2-oxepanone 5-Ethyl-2-oxepanone cis-3,7-Dimethyl-2oxepanone

5. After about 1 h of induction, 2-methyl cyclohexanone (20.0 g) was dispersed into the cell medium for the biotransformation. 6. Samples of 10 mL were withdrawn from the fermenter during the course of biotransformation. Sample (1 mL) was immediately extracted with an equal volume of ethyl acetate. The extracted sample solutions were analyzed by GC. Sample (1 mL) was also extracted with dodecane (0.2 mL) for fast determination of the biotransformation using a ReactIR 4000. The cell density was measured using a spectrophotometer and the residual glucose in the aqueous phase was monitored using high-performance liquid chromatography. 7. Downstream extraction. The culture broth was diluted with ethyl acetate and the aqueous phase separated using a separation funnel. The organic layer was collected and dried over anhydrous sodium sulfate. Removal of the solvent by rotary evaporator gave (S)-7-methyl-2-oxepanone as a light yellow oil (6.5 g, 38 % yield). Chiral-phase GC showed 99 % ee and >97 % purity. EI-MS and NMR confirmed the product. Note: the unconverted (R)-2-methyl cyclohexanone evaporated completely under the aeration conditions used during the overnight incubation. 11.8.5

Conclusion

CPDMO is a new bioreagent for the synthesis of optically pure lactones with excellent enantioselectivity. CPDMO is not only effective in desymmetrization of meso and prochiral compounds (Procedure 2, Section 11.8.2), but excellent in carrying out the kinetic resolution of racemates (Procedure 3, Section 11.8.3). Additional examples of optically pure lactones that can be obtained are summarized in Table 11.4. In the fermenter work (Procedure 4, Section 11.8.4), (R)–2-methyl cyclohexanone was not converted, but evaporated under aeration condition (1 vvm). This led to the expected product (S)-7-methyl oxepanone at the end of the experiment. The optically pure lactone could be recovered without silica-gel chromatography separation. However, the production yield may be improved by using a better condenser.

References 1. Stewart, J.D., Cyclohexanone monooxygenase: a useful reagent for asymmetric Baeyer–Villiger reactions. Curr. Org. Chem., 1998, 2, 195–216. 2. Iwaki, H., Hasegawa, Y., Wang, S., Kayser, M.M. and Lau, P.C.K., 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 1,2-monooxygenase. Appl. Environ. Microbiol., 2002, 68, 5671-5684.

350


3. Mihovilovic, M.D., Rudroff, F. and Grotzl, B., Enantioselective Baeyer–Villiger oxidations. Curr. Org. Chem., 2004, 8, 1057–1069. 4. Ten Brink, G.-J., Arends, I.W.C.E. and Sheldon, R.A., The Baeyer–Villiger reaction: towards greener procedures. Chem Rev., 2004, 104, 4105–4123. 5. Constable, D.J.C., Dunn, P.J., Hayler, J.D., Humphrey, G.R., Leazer, Jr, J.L., Linderman, R.J., Lorenz, K., Manley, J., Pearlman, B.A., Wells, A., Zaks, A. and Zhang, T.Y., Key green chemistry research areas – a perspective from pharmaceutical manufacturers. Green Chem., 2007, 9, 411–420. 6. Iwaki, H., Wang, S., Grosse, S, Bergeron, H., Nagahashi, A., Lertvorachon, J., Yang, J., Konishi, Y., Hasegawa, Y. and Lau, P.C.K., Pseudomonad cyclopentadecanone monooxygenase displaying an uncommon spectrum of Baeyer-Villiger oxidations of cyclic ketones. Appl. Environ. Microbiol., 2006, 72, 2707–2720. 7. Sambrook, J.E., Fritsch E.F. and Maniatis, T. Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press: Cold Spring Harbor, NY, 1989. 8. Panke, S., Held, M., Wubbolts, M.G., Witholt, B. and Schmid, A., Pilot-scale production of (S) styrene oxide from styrene by recombinant Escherichia coli synthesizing styrene monooxygenase. Biotechnol. Bioeng., 2002, 80, 33–41.

12 Whole-cell Oxidations and Dehalogenations

12.1

Biotransf ormations of Naphthalene to 4-Hydroxy-1-tetralone by Streptomyces griseus NRRL 8090 Arshdeep Khare, Andrew S. Lamm and John P.N. Rosazza

Streptomyces griseus NRRL 8090 catalyzes a series of biotransformations of naphthalene and 2-methyl-1,4-naphthaquinone to their corresponding racemic and diastereomeric 4-hydroxy-1-tetralones (Figure 12.1). The yields of 4-hydroxy-1tetralone obtained with S. griseus are much higher than those produced by various fungi that oxidize naphthalene.1 12.1.1 12.1.1.1 • • • • • • • •

Procedure 1: Cultivation of S. griseus NRRL 8090 Materials and Equipment

Glycerol (20 g) soybean flour (30 g) sterile water (5 mL) culture of S. griseus NRRL 8090 stored on Sabouraud maltose agar slant at 4 C distilled water sterile loop two 125 mL DeLong culture flasks with stainless steel cap rotary shaker.



352

Whole-cell Oxidations and Dehalogenations OH

Naphthalene

O

O

1-Naphthol

OH 4-Hydroxy-1-Tetralone

1-Tetralone

O

O

CH3

CH3

OH 2-Methyl-4-Hydroxy-1-Tetralone

O 2-Methyl-1,4-Naphthoquinone

Figure 12.1 S. naphthaquinone

12.1.1.2

griseus-catalyzed

oxidation

of

naphthalene

and

2-methyl-1,4-

Procedure

1. Cultures were grown in a two-stage procedure in 25 mL of soybean flour and glycerol medium (30 g soybean flour and 20 g glycerol in 1 L distilled water) held in stainlesssteel capped, 125 mL DeLong culture flasks. The flasks containing the medium were autoclaved at 15 psi at 121 C for 15 min. The surface growth from slants was suspended in 5 mL of sterile water with a sterile loop and used to inoculate 25 mL sterile medium (Stage I culture). Cultures were incubated for 72 h, at 29 C, with shaking at 200 rpm. A 10 % inoculum derived from the 72-h-old Stage I culture was used to inoculate sterile medium (Stage II culture), which was incubated for 24 h before adding naphthalene substrate for biotransformation. 12.1.2

Procedure 2: Synthesis of 4-Hydroxy-1-tetralone O

OH

12.1.2.1 • • • • •


Distilled water naphthalene (150 mg) N,N-dimethylformamide (DMF, 30–40 mL) ethyl acetate hexanes

12.1 Biotransformations of Naphthalene to 4-Hydroxy-1-tetralone

• • • • • • • • •

353

silica gel (GF254) plates, 0.25 mM thin-layer chromatography (TLC) solvent system: ethyl acetate:hexanes (50:50 v/v) 254 nm UV lamp for TLC plate visualization anhydrous sodium sulfate ten 125 mL DeLong flasks with stainless-steel caps Shimadzu GC-17A series RTX-5 column, 15 m (length), 0.25 mm (i.d.) and 0.15 m film thickness. rotary evaporator desktop centrifuge.

12.1.2.2

Procedure

2. For analytical purposes, Stage II cultures of S. griseus were prepared in 125 mL DeLong culture flasks as described. The cultures were shaken at 200 rpm at 29 C for 24 h and then 10 mg of substrate naphthalene in 30–40 mL of DMF was added to each 25 mL volume of culture and incubations were continued with shaking. 3. Samples (3 mL) of substrate containing cultures were taken at 24, 48 and 120 h after substrate addition and extracted with equal volumes of ethyl acetate. The organic phases were separated by centrifugation for 3 min in a desktop centrifuge and used for TLC analysis. 30–40 mL of sample extracts were spotted onto TLC plates that were developed with ethyl acetate:hexane (v/v). Visualization of TLC plates was done by fluorescence quenching under 254 nm UV light. Rf values were: naphthalene, 0.90; 1-naphthol, 0.85; 1-tetralone, 0.8; 4-hydroxy-1-tetralone, 0.32; menadione, 0.86; and 2-methyl-4-hydroxy-1-tetralone, 0.43. 4. Preparative-scale biotransformation of 150 mg naphthalene was conducted using ten 125 mL DeLong flasks, each containing 25 mL of 24-h-old Stage II cultures and 15 mg of naphthalene in DMF (30–40 mL). After 120 h, contents of all flasks were combined, centrifuged at 7000g for 20 min. The supernatant was extracted three times with 150 mL ethyl acetate and cells washed twice with 20 mL of ethyl acetate each time. Organic extracts were combined, washed with distilled H2O, dried over anhydrous Na2SO4 and concentrated in vacuo. The residue was dissolved in a minimum amount of ethyl acetate, applied to a 2 22 cm silica-gel column and eluted with a hexane ethyl acetate gradient ranging from 100:3 to 75:25. 4-Hydroxy-1-tetralone was obtained in 43 % yield. 1

H NMR (CDCl3, 400 MHz) 2.17 (1H, m, H-3), 2.41 (1H, m, H-3), 2.58 (1H, ddd, J ¼ 17.8, 9.6, 4.8 Hz, H-2), 2.92 (1H, ddd, J ¼ 17.8, 7.5, 4.6 Hz, H-2), 4.98 (1H, dd, J ¼ 8.1, 3.9 Hz, H-4), 7.41 (1H, m, H-7), 7.60 (2H, m, H-5 and H-6), 8.03 (1H, d, J ¼ 7.7 Hz, H-8). 12.1.3

Procedure 3: Synthesis of 2-Methyl-4-hydroxy-1-tetralone O Me

OH

354

Whole-cell Oxidations and Dehalogenations

1. The same method described in Procedure 2 (Section 12.1.2) was used for preparative-scale biotransformation of 150 mg of 2-methyl-1,4-naphthoquinone, except that reactions were incubated for only 72 h before being combined, centrifuged, extracted and chromatographically purified to give 50 % yield (92 mg) of product. 1

H NMR (CDCl3, 400 MHz) 1.15 (CH3, 3H, d, J ¼ 6.8 Hz), 1.30 (CH3, 3H, d, J ¼ 6.6 Hz), 1.42 (CH3, 3H, d, J ¼ 6.6 Hz), 2.51 (1H, m, H-3), 2.60 (1H, m, H-3), 2.8 (1H, m, H-2), 5.04 (1H, dd, J ¼ 11.1, 4.8 Hz, H-4), 7.38 (1H, m, H-7), 7.58 (1H, m, H-6), 7.7 (1H, d, J ¼ 7.52 Hz, H-5), 8.02 (1H, m, H-8); 13 C NMR (CDCl3, 100 MHz), 16.31, 16.43, and 17.21 (3-CH3); and signals for three 199.20, 200.67, 201.50 (3-C¼O).

References and Notes 1. Gopishetty, S.R., Heinemann, J., Deshpande, M. and Rosazza, J.P.N., Aromatic oxidations by Streptomyces griseus: biotransformations of naphthalene to 4-hydroxy-1-tetralone. Enzyme Microbiol Technol., 2007, 40, 1622. 2. For gas chromatography analysis, samples were spiked with 2-methyl-naphthalene as an internal standard. Samples were analyzed using a Shimadzu GC-17A series gas chromatograph equipped with RTX-5 column, 15 m (length) 0.25 mm (i.d.) and 0.25 mm (film thickness). The initial column temperature was 70 C and temperature was increased at 20 C min1 300 C, and column temperature was held for 13 min. Retention times Rt: naphthalene, 3.2 min; 2-methylnaphthalene, internal standard, 4.09 min; 1-tetralone, 4.7 min; menadione, 5.68 min; 1-naphthol, 5.7 min; 4-hydroxy-1-tetralone, 6.1 min; and 2-methyl-4-hydroxy-1-tetralone, 6.18, 6.27, 6.3 and 6.4 min.

12.2 Hydroxylation of Imidacloprid for the Synthesis of Olefin Imidacloprid

12.2

355

Hydroxylation of Imidacloprid for the Synthesis of Olefin Imidacloprid by Stenotrophomonas maltophilia CGMCC 1.1788 Sheng Yuan and Yi-jun Dai

Resting cells of bacterium Stenotrophomonas maltophilia CGMCC 1.1788 catalyze the stereoselective hydroxylation at position C12 of imidacloprid (IMI) in the imidazolidine ring to form (R)-5-hydroxy IMI. Under acidic conditions, 5-hydroxy IMI is converted into olefin IMI (Figure 12.2), which exhibits about 19 times more insecticidal efficacy than IMI against horsebean aphid imago. 12.2.1 12.2.1.1 • • • • • • • • • • •

Procedure 1: Cultivation of S. maltophilia CGMCC 1.1788 Materials and Equipment

Luria–Bertani (LB) broth tryptone (35 g) yeast extract (17.5 g) NaCl (35 g) distilled water (3.5 L) stored culture of S. maltophilia CGMCC 1.1788 one plate, 11 cm flask with a poromeric silicone plug, 1 L fermentor, 5 L shaker high-speed freeze centrifuge.

12.2.1.2

Procedure

1. A single colony of bacterium S. maltophilia CGMCC 1.1788 strain on LB agar plate is inoculated to a 1 L flask containing 300 mL of LB broth and cultivated in a rotary shaker at 220 rpm at 30 C for 13 h. Then, the culture broth is poured into the fermentor containing 3.2 L LB broth for cultivation. During cultivation, the fermentor is constantly aerated and stirred at 500 rpm at 30 C. After cultivation for 10 h, the fermentation broth is centrifuged at 6000g for 20 min to obtain the cells of S. maltophilia CGMCC 1.1788 (about 60 g wet weight).

Figure 12.2 Chemical structure of IMI and its transformation products

356


12.2.2

12.2.2.1 • • • • • • • • • • • • • • • •

Procedure 2: Synthesis of 5-Hydroxy IMI


KH2PO4 (1.3609 g) Na2HPO412H2O (68.0466 g) IMI (3.0 g) sucrose (150 g) distilled water (3 L) anhydrous sodium sulfate (30 g) dichloromethane (6.1 L) ethyl acetate (3 L) acetonitrile (10 mL) ultrafiltration membranes, 0.22 mm pore size one flask, 5 L beaker, 50 mL one separatory funnel, 12 L fermentor, 5 L rotary evaporator vacuum pump

12.2.2.2

Procedure

1. Fresh harvested cells were suspended in 3.0 L of 87 mmol L1 phosphate buffer (pH 8.0) with 3 g IMI and 150 g sucrose in a 5 L fermentor for transformation. During transformation, the fermentor was constantly aerated and stirred at 500 rpm at 30 C for 72 h. At the end of transformation, cells were removed by centrifugation at 6000g for 20 min and the supernatant is collected. 2. The supernatant was first extracted with dichloromethane (2 3 L) to eliminate the remaining IMI. The aqueous fraction was then extracted with ethyl acetate (3 L). The ethyl acetate extract, containing 5-hydroxy IMI, wais dried with 30 g anhydrous sodium sulfate and concentrated to about 1/20th of the original volume in a vacuum rotary evaporator and then filtered with 0.22 mm pore size ultrafiltration membranes. The filtered solution was evaporated again until white crystals were produced. The crystals were filtered, washed twice with dichloromethane and then dissolved in 10 mL acetonitrile by heating. At 4 C, the 5-hydroxy IMI crystallized from the above solution and was filtered and dried under vacuum. A total of 413 mg of 5-hydroxy IMI was obtained.

12.2 Hydroxylation of Imidacloprid for the Synthesis of Olefin Imidacloprid

357

1

H NMR (dimethylsulfoxide (DMSO); 400 MHz) 9.17 (s, 1H, H-10), 8.38 (d, J ¼ 2.4 Hz, 1H, H-2), 7.82 (dd, J ¼ 8.2, 2.4 Hz, 1H, H-4), 7.51 (d, J ¼ 8.2 Hz, 1H, H-5), 6.82 (d, J ¼ 7.5 Hz, 1H, 12-OH), 5.25 (ddd, J ¼ 7.5, 7.5, 2.5 Hz, 1H, H-12), 4.58 (d, J ¼ 16.1 Hz, 1H, H-7), 4.42 (d, J ¼ 16.1 Hz, 1H, H-7), 3.84 (dd, J ¼ 12.0, 7.6 Hz, 1H, H-11), 3.37 (J ¼ 12.0, 2.4 Hz, 1H, H-11). 13C NMR (DMSO; 100 MHz) 158.9 (C9), 149.7 (C2), 149.6 (C6), 139.7 (C4), 132.9 (C3), 124.5 (C5), 80.3 (C12), 50.6 (C11), 41.8 (C7). 12.2.3

12.2.3.1 • • • • • • • • •

Procedure 3: Synthesis of Olefin IMI


Distilled water (350 mL) hydrochloric acid (5 mL) 5-hydroxyl IMI (0.3 g) ethyl acetate (100 mL) anhydrous sodium sulfate (5 g) one beaker, 1 L one separatory funnel, 1 L vacuum rotary evaporator water bath.

12.2.3.2

Procedure

1. 5-Hydroxyl IMI (0.3 g) was added to 350 mL distilled water and heated to 80 C to obtain a solution. Hydrochloric acid (5 mL) was added and the solution heated at 80 C for 35 min. After cooling to room temperature, the reaction solution was extracted with ethyl acetate (350 mL). The extracts were dried with 5 g anhydrous sodium sulfate and concentrated in a vacuum rotary evaporator until the product appeared as white needle crystals. The crystals were collected and dried in air (0.1 g). 1

H NMR (DMSO; 400 MHz) 12.79 (s, 1H, H-10), 8.41 (s, 1H, H-2), 7.77 (d, J ¼ 8.0 Hz, 1H, H-4), 7.53(d, J ¼ 8.0 Hz, 1H, H-5), 7.38 (s, 1H, H-12), 7.07(s, 1H, H-11), 5.13 (s, 2H, H-7). 13C NMR (DMSO; 100 MHz) 150.3 (C9), 149.8 (C2), 146.2 (C6), 139.8 (C4), 131.8 (C3), 124.9 (C5), 117.3 (C12), 114.3 (C11), 45.1 (C7). 12.2.4

Conclusion

The procedure is very easy to reproduce and the stereoselective hydroxylation of IMI with S. maltophilia CGMCC 1.1788 may be applied to some other neonicotinoid insecticides, such as thiacloprid (Table 12.1).

358

Whole-cell Oxidations and Dehalogenations Table 12.1 Transformation of substrates by S. maltophilia CGMCC 1.1788 Substrates

Products

Transformation yield (%)

26

28

23

References 1. Dai, Y.J., Yuan, S., Ge, F., Chen, T., Xu, S.C. and Ni, J.P., Microbial imidacloprid for the synthesis of highly insecticidal olefin imidacloprid. Biotechnol., 2006, 71, 927–934. 2. Dai, Y.J., Chen, T., Ge, F., Huan, Y., Yuan, S. and Zhu, F.F., Enhanced imidacloprid by Stenotrophomonas maltophilia upon addition of sucrose. Biotechnol., 2007, 74, 995–1000.

hydroxylation of Appl. Microbiol. hydroxylation of Appl. Microbiol.

12.3 Biocatalytic Synthesis of 6-Hydroxy Fluvastatin

12.3

359

Biocatalytic Synthesis of 6-Hydroxy Fluvastatin using Mortierella rammaniana DSM 62752 in Shake Flask Culture and on Multi-gram Scale using a Wave Bioreactor Matthias Kittelmann, Maria Serrano Correia, Anton Kuhn, Serge Parel, Ju¨rgen Ku¨hno¨l, Reiner Aichholz, Monique Ponelle and Oreste Ghisalba

Fluvastatin is a serum cholesterol-lowering drug belonging to the class of ‘statins’, which acts through inhibition of 3-hydroxy-3-methyl-glutaryl coenzyme A (HMG-CoA) reductase, the rate-limiting enzyme in cholesterol biosynthesis.1 The 5- and 6-hydroxy and the N-de-isopropyl derivative represent the major human metabolites of this drug.2 The synthesis of oxidized drug metabolites via microbial biotransformation has broadly been discussed in the literature in recent years.3–5 We evaluated the biotransformation of fluvastatin using different bacterial and fungal wild-type strains as an alternative to chemical synthesis. With Mortierella (M.) rammaniana DSM 62752 6-hydroxy fluvastatin was produced (Figure 12.3) in multi-hundred milligram amounts via shake flask culture and in gram amounts using a BioWave bioreactor. 5-Hydroxy fluvastatin was synthesized with Streptomyces violascens ATCC 31560 on multi-milligram scale, though with much lower yield, so that this method will not be outlined in detail. 12.3.1 12.3.1.1 • • • • • • • • • • •

Procedure 1: Reactivation of M. rammaniana DSM 62752 from a Frozen Culture on Agar Plates Materials and Equipment

Malt extract (2.25 g) casitone (0.375 g) agar (1.13 g) distilled water (75 mL) culture of M. rammaniana DSM 62752 frozen at 80 C glass bottle, 200 mL, screw capped three Petri dishes inoculation loop, sterile steam-sterilizator water bath, temperature controlled incubator, temperature controlled.

12.3.1.2

Procedure

1. Malt extract (2.25 g), casitone (0.375 g) and agar (1.13 g) were dissolved in 75 mL of distilled water in, for example, a 200 mL screw-capped glass bottle. The screw cap was not completely closed and the mixture together with a magnetic bar sterilized in the steam-sterilizer for 20 min at 121 C. 2. The hot liquid agar medium was mixed by magnetic stirring and cooled to 45 C in a temperature-controlled water bath. Then the agar was poured into three Petri dishes and solidified by cooling to room temperature. 3. The agar plates were inoculated on the whole surface from a culture of M. rammaniana frozen at 80 C using a sterile inoculation loop and incubated for 4 days at 28 C.

360

Whole-cell Oxidations and Dehalogenations F

Mortierella rammaniana DSM 62572

F

6 HO

O

N OH

OH

Na+

O

6-Hydroxy fluvastatin-Na O

N OH

OH

O

Na+

F

Streptomyces violascens ATCC 31560

Fluvastatin-Na

HO 5 O

N OH

OH

O

Na+

5-Hydroxy fluvastatin-Na

Figure 12.3

12.3.2

Synthesis of 5- and 6-hydroxy fluvastatin by microbial biotransformation

Procedure 2: Preculture and Main Culture of M. rammaniana and Synthesis of 6-Hydroxyfluvastatin F

HO

OH

N OH

12.3.2.1 • • • • • • • • • • • • • • • •

OH

O


Distilled water (16.5 L) glucose (281 g) Lab-Lemco (Oxoid) (42 g) peptone from casein (52.5 g) yeast extract (52.5 g) casitone (Becton Dickinson) (31.5 g) NaCl (15.75 g) 3-morpholino propane sulfonic acid (MOPS) (220.5 g) NaOH solution, 4 M fluvastatin-Na (1 g, 2.23 mmol) methanol (10 mL) XAD-16 adsorber resin (16 g) (Rohm and Haas France S.A.S., Lauterbourg, France) isopropanol (2 L) ethyl acetate (4.4 L) saturated NaCl solution (800 mL) NaCl (100 g)


361

• MgSO4, anhydrous • RP18 silica gel (30 g) (Lichroprep RP18 40–60 mm, Merck KGaA, Darmstadt, Germany) • acetonitrile, high-performance liquid chromatography (HPLC) gradient grade (280 mL) • KH2PO4 (0.134 g, 0.98 mmol) • Na2SO4, anhydrous • 25 Erlenmeyer flasks, 2 L, four baffles • 5 Erlenmeyer flasks, 500 mL, one baffle • 1 Erlenmeyer flask, 1 L • cotton • gauze • inoculation loop, sterile • steam sterilizer • laboratory shaker, 5 cm agitation radius • 20–30 pipettes, 25 mL, sterile • 10–15 pipettes, 1 mL, sterile • polypropylene tube, 50 mL, screw capped, presterilized (e.g. Falcon tubes, Becton Dickinson Labware, Franklin Lakes, NJ, USA) • filter funnel • sinter glass filter funnel • separatory funnel for 1 L extraction • filter paper • rotary evaporator • high-vacuum pump. 12.3.2.2

Procedure

Growth of M. rammaniana and Biotransformation of Fluvastatin 1. Glucose (281 g), Lab-Lemco (Oxoid) (42 g), peptone from casein (52.5 g), yeast extract (52.5 g), casitone (31.5 g), NaCl (15.75 g), and MOPS (220.5 g) were dissolved in 10.5 L of distilled water and the pH was adjusted to 6.5 with 4 M NaOH. 2. The resulting solution was filled in 400 mL portions into 25 Erlenmeyer flasks with 2 L total volume equipped with four baffles and in 100 mL portions in five Erlenmeyer flasks with 500 mL total volume equipped with one baffle. The flasks were closed by cotton plugs wrapped in gauze and autoclaved at 121 C for 20 min. 3. The five flasks with each 100 mL of medium (precultures) were inoculated with mycelium of M. rammaniana from the agar plates using a sterile inoculation loop and incubated on a laboratory shaker with 5 cm agitation radius at 28 C and 220 rpm for 3 days. 4. Each of the 2 L Erlenmeyer flasks (main cultures) was then inoculated with 20 mL of preculture and incubated at 28 C and 180 rpm. 5. 1 g of fluvastatin-Na was dissolved in 10 mL of methanol (fluvastatin solution) in, for example, a presterilized, screw-capped 50 mL polypropylene tube. Glucose (50 g) was dissolved 0.5 L of distilled water and sterilized at 121 C for 20 min.

362


6. After 48 h of incubation, 0.4 mL of the methanolic fluvastatin solution and 20 mL of the sterile glucose solution were added to each of the 2 L shake flasks under sterile conditions. Incubation under shaking was continued for another 42 h. The degree of conversion was measured by analytical RP18-HPLC with diode array detection.6 Purification of 6-Hydroxy Fluvastatin 1. To each of the flasks, 16 g of the adsorber resin XAD-16 was added and the flasks were shaken for a further 3 h. The resin was collected by filtering off over gauze in a filter funnel and washed with 4 L of distilled water. Then it was eluted four times with portions of 500 mL of isopropanol by gentle shaking in a 1 L Erlenmeyer flask for 30 min and filtering off the resin. The solvent was removed under reduced pressure at 30 C bath temperature. The residue was dissolved in 400 mL of ethyl acetate and washed twice with 400 mL of saturated NaCl solution. The organic phase was dried over anhydrous MgSO4 and the solvent removed under reduced pressure at 30 C bath temperature. Further purification was performed via a second solid-phase extraction on RP18 silica gel. 2. The crude extract (3 g) was dissolved in acetonitrile (30 mL) and mixed with dry solid RP18-phase (30 g) and 270 mL of potassium phosphate buffer 0.7 mM pH 7 ( ¼ Kpibuffer, preparation: KH2PO4 (0.134 g, 0.98 mmol) dissolved in 1400 mL distilled water, pH adjusted to 7 with 0.1 M KOH). The mixture was filtered in a sinter-glass filter funnel and the RP18 silica gel was washed with a 10 % (v/v) solution of acetonitrile in Kpi-buffer (300 mL). Subsequently, the RP18 silica gel was eluted twice with 500 mL of a 25 % (v/v) solution of acetonitrile in Kpi-buffer. To each of the two resulting fractions, 50 g of NaCl was added and they were extracted twice with 500 mL of ethyl acetate. The two organic phases were dried over anhydrous Na2SO4, filtered over filter paper and the solvent was removed under reduced pressure at 20 C and finally under high vacuum for 2 h. (Fraction 1: 480 mg, light brown resin, 62 % purity RP18 HPLC-UV205 nm, 57 % RP18 HPLC–mass spectrometry (MS), >27 % molar yield, structure identification in comparison with chemically synthesized 6-hydroxy fluvastatin). 1

H NMR (400 MHz, dimethylsulfoxide) ¼ 1.19 (6H, dd, appears as t), 1.29 (1H, m), 4.83 (1H, hept J ¼ 8 Hz), 5.66 (1H, dd, J ¼ 4 and 16 Hz), 6.50–6.53 (2H, m), 7.20–7.29 (4H, m), 7.40–7.45 (2H, m). 12.3.3 12.3.3.1 • • • • •

Procedure 3: Synthesis of 6-Hydroxy Fluvastatin with M. rammaniana DSM 62752 in a BioWave Bioreactor on 22 L Scale Materials and Equipment

Distilled water 23 L Tween 80 (70 mg) glycerol (14 g) cellulose powder (4 g) oat meal (2 g)


363

• tomato paste (2 g), low salt, no preservatives, in the original recipe the brand is Hunt’s Tomato Paste • KH2PO4 (11.3 g) • MgSO4 (0.2 g) • agar (4 g) • glucose (44 g) • malt extract (220 g) • yeast extract (88 g) • NH4Cl (11 g) • 2-morpholino ethane sulfonic acid monohydrate (MES, 429 g) • antifoam 204 (Sigma) • antifoam Y-30 solution (Sigma) • glass bottle, screw capped, 100 mL • glass bottle, screw capped, 500 mL • magnetic bar • steam sterilizator • water bath, temperature controlled • eight Petri dishes • 10 pipettes, 10 ml, sterile • 10 Eppendorf tips, truncated, sterile • 10 L-shaped spreaders, plastic, sterile (e.g. VWRI612-1560, VWR International) • two polypropylene tube, 50 mL, screw-capped, presterilized (e.g. Falcon tubes, Becton Dickinson Labware, Franklin Lakes, NJ, USA) • BioWave 50SPS bioreactor (Wave Biotech AG, Tagelswangen, Switzerland) (since recently distributed by Sartorius BBI Systems GmbH, Melsungen, Germany, as ‘Biostat Cultibag RM 50’) • Wavebag 50 L total volume, exhaust gas line 0.5 inch • peristaltic pump (Heidolph Pumpdrive 5006) • sterile microfiltration capsule, pore size 0.45 þ 0.2 mm (Sartobran 300, 5231307-H5–00, Sartorius Biotech GmbH, Go¨ttingen, Germany) • six sterile disposable syringes, 50 mL • 50–60 sterile disposable syringes, 10 mL (for sampling) • membrane pump KNF Labport type N86KN.18 (KNF Neuberger, Freiburg, Germany) • thermal mass flowmeter for air, type GCA-B5SA-BA20 and • thermal mass flowmeter for oxygen, type GCR-A9SA-BA15 (Thermal Mass Flow Co., USA) • sterilized glass bottle, 1 L, screw capped (as foam trap) • spectrophotometer. 12.3.3.2

Procedure

Preparation of Spore Suspension 1. Tween–glycerol solution. Tween 80 (70 mg) and glycerol (14 g) were dissolved in distilled water (59 mL) and sterilized at 121 C for 20 min.

364


2. Preparation of sporulation agar plates. Cellulose powder (4 g), oat meal (2 g), tomato paste (2 g), KH2PO4 (0.3 g), MgSO4 (0.2 g) and agar (4 g) were dissolved in 200 mL of distilled water in a 500 mL screw-capped glass bottle; the screw cap was not completely closed, and the mixture together with a magnetic bar was sterilized in the steamsterilizator for 20 min at 121 C; the hot liquid agar medium was mixed by magnetic stirring and cooled to 45 C in a temperature-controlled water bath; then the agar was filled into eight Petri dishes and solidified by cooling to room temperature. 3. 7 mL of Tween–glycerol solution was mixed with M. rammaniana mycelium grown densely on a malt extract agar plate (see Procedure 1, Section 12.3.1) using a sterile plastic L-shaped spreader. 4. Portions (100 mL) of mycelium suspension were transferred to eight sporulation agar plates with a sterile, truncated Eppendorf tip and spread with a sterile plastic L-shaped spreader. 5. The plates were incubated for 8 days at 28 C. Then Tween–glycerol solution (7 mL) was filled onto each plate and the spores/mycelium were suspended by rigorous scraping of the agar surface again with a sterile plastic L-shaped spreader. The spore/mycelium suspension was stored in two sterile 50 mL polypropylene tubes at 80 C until use. Growth of M. rammaniana and Biotransformation In BioWave bioreactors, a disposable polyethylene bag (Wavebag) serves as the cell containment which is rocked on a temperature-controlled table for mixing and gas exchange. Oxygen is supplied via a stream of sterile air/oxygen mixture through the headspace of the bag. For the cultivation of highly oxygen-demanding microorganisms the required gas flow exceeds the capacity of the built-in pump, so that an external membrane pump had to be employed. Furthermore, supplementation with pure oxygen was necessary. 1. Preparation of concentrated medium. Glucose (44 g), malt extract (220 g), yeast extract (88 g), KH2PO4 (11 g), NH4Cl (11 g) and MES (429 g) was dissolved in 4.4 L of distilled water, the pH was adjusted to 5.9 with concentrated NaOH and the liquid was sterilized at 121 C for 30 min in a steam sterilizer. 2. The Wavebag (50 L volume) was placed on the temperature-controlled tray and completely inflated with air using a membrane pump. The airflow was adjusted to 2250 mL using a thermal mass flow meter. 3. The concentrated sterilized medium was pumped into the Wavebag with a peristaltic pump through autoclaved silicone tubes. Then distilled water (17.6 L) was pumped in through a disposable, sterile microfiltration capsule. Rocking was started at an angle of 10.5 at 36–37 rpm as well as the temperature regulation, set-point 28 C. 4. When the temperature was equilibrated, spore suspension (28 mL) was added with a sterile disposable 50 mL syringe through the sampling port, followed by 10 mL of a heat-sterilized antifoam 204 emulsion (1 mL antifoam 204 mixed with 9 mL of distilled water). Then, the supply of pure oxygen was started and adjusted to 250 mL min1 using a second thermal mass flow meter. The mixing of air and oxygen in the desired ratio was effected by joining the line for pure oxygen and the gas inlet of the pump (aspiration port) with a ‘T’ connector, allowing both gases to be taken in (Figure 12.4). In the gas inline tubing between the air flow meter and the Wavebag, a hydrostatic pressure relief valve was built in, providing a maximum backpressure of 20 cm water


Air

Oxygen

Overpressure release via 20 cm watercolumn

Membrane Air pump flow meter F

F

365

Sterile inlet filter

1/2" exhaust tubing

Oxygen flow meter

Exhaust air Sterile filter

Wavebag Foam trap

Figure 12.4 Gas flow in the BioWave reactor under oxygen supplementation

column. In the exhaust gas line between the Wavebag and the sterile filter, a sterile 1 L screw-capped glass bottle containing a few millilitres of antifoam Y-30 emulsion was included as a foam trap. 5. During the day, samples were taken around every 2 h and analysed without dilution for pH. Growth was estimated by measuring the optical density at 600 nm (OD600) against distilled water in samples diluted to OD600 £ 0.3 with distilled water using a spectrophotometer. 6. After 30 h from inoculation, NaOH solution (4 m, 40 ml) was injected into the Wavebag with a sterile disposable syringe. 8.8 g of fluvastatin-Na was dissolved in 88 mL of methanol. At 48, 74, 120, and 192 h after inoculation with spores, fluvastatin solution (22 mL) was supplemented to the culture via the sampling port and a sterile disposable 50 mL syringe. The degree of conversion was measured by analytical RP18-HPLC with diode array detection.6 7. After 10–12 days, the maximum product concentration was achieved and fluvastatin completely consumed. The 6-hydroxy-fluvastatin-containing culture liquid was stored in the Wavebag at 20 C for later purification. In other cases, the culture liquid was conveniently and safely harvested from the Wavebag without aerosol formation by sucking into glass bottles in a vacuum line with a sterile filter installed between the collecting vessel and the vacuum pump. 12.3.4

Conclusion

The synthesis of 6-hydroxy fluvastatin with M. rammaniana DSM 62752 gave high conversion (>95 %) in shake flask culture on 400 mL scale with 0.1 g L1 of fluvastatin as well as on 22 L scale in a Wave bioreactor-fed batch process at a final substrate concentration of 0.4 g L1. Instead of the partial purification by a second solid-phase extraction described above, 6-hydroxy fluvastatin can be obtained in high purity (95 %) by, for example, preparative medium-pressure liquid chromatography (MPLC) on RP18 silica gel.7 5-Hydroxy fluvastatin could be prepared analogously via biotransformation in shake flask culture with Streptomyces violascens ATCC 31560. Different media and minor variations of the process schedule had to be applied.8 Before supplementation of

366


fluvastatin it was important that glucose had been completely consumed (check with urine–glucose test sticks). Furthermore, the pH had to be stabilized in the culture by addition of CaCO3 at the time of fluvastatin addition.9 Since the organism produced both 5- and 6-hydroxy fluvastatin, purification via RP18-LC was needed.7

References and Notes 1. Christians, U., Jacobsen, W. and Floren, L.C., Metabolism and drug interactions of 3-hydroxy-3methylglutaryl coenzyme A reductase inhibitors in transplant patients: are the statins mechanistically similar? Pharmacol. Ther., 1998, 80, 1–34. 2. Fischer, V., Johanson, L., Heitz, F., Tullmann, R., Graham, E., Baldeck, J.P. and Robinson, W.T., The 3-hydroxy-3-methylglutaryl coenzyme A reductase inhibitor fluvastatin: effect on human cytochrome P-450 and implications for metabolic drug interactions. Drug Metab. Dispos., 1999, 27, 410–416. 3. Azerad, R., Microbial models for drug metabolism. In Biotransformations, Faber, K. and Scheper, T. (eds), Advances in Biochemical Engineering/Biotechnology, vol. 63, Springer, 1999, pp. 169–218. 4. Venisetty, R.K. and Ciddi, V., Application of microbial biotransformation for the new drug discovery using natural drugs as substrates. Curr. Pharm. Biotechnol., 2003, 4, 153–167. 5. Ghisalba, O. and Kittelmann, M., Preparation of drug metabolites using fungal and bacterial strains. In Modern Biooxidation – Enzymes, Reactions and Applications, Schmid, R.D. and Urlacher, V.B. (eds). Wiley–VCH Verlag, Weinheim, 2007, pp. 211–232. 6. Sample preparation. Culture liquid (0.5 mL) was mixed with isopropanol (0.5 mL), kept at room temperature for 10 min and centrifuged at 25 000g and 20 C in a refrigerated Eppendorf centrifuge. The supernatant was subjected to HPLC-analysis. HPLC system Agilent 1100; column: Chromolith Performance RP-18e 100 mm 4.6 mm, pre-column Chromolith Guard Cartridge RP-18e 5 mm 4.6 mm (Merck KgaA, Darmstatt, Germany); elution: flow rate 2 mL min1, eluent A ¼ 3 mM H3PO4, eluent B ¼ acetonitrile (gradient grade), gradient 5–100 % B in 4.75 min; injection volume 10 mL; diode array detection 190–400 nm. 7. Preparative MPLC. Solid phase Lichroprep RP18 40–60 mm (Merck KGaA, Darmstadt, Germany), first gradient 5–45 % acetonitrile against 1 mM ammonium formate (six column volumes), second gradient 5–50 % acetonitrile with 1 mM formic acid as the aqueous phase. 8. Variations for 5-hydroxy fluvastatin. Medium for growth on agar plates: Plate Count Agar (Fluka/ Sigma Aldrich, Buchs, Switzerland); medium for preculture and main culture: glucose 20 g L1, soytone (Becton Dickinson) 15 g L1, yeast extract 10 g L1, pH adjusted to 6.5 with NaOH. Incubation time of main culture before fluvastatin addition 3 days; biotransformation period 24 h. 9. Each flask (400 mL culture) was supplemented with a steam-sterilized (121 C) suspension of CaCO3 (3 g) in distilled water (30 mL) immediately after fluvastatin addition.

12.4 Synthesis of 1-Adamantanol from Adamantane

12.4

367

Synthesis of 1-Adamantanol from Adamantane through Regioselective Hydroxylation by Streptomyces griseoplanus Cells Koichi Mitsukura,* Yoshinori Kondo, Toyokazu Yoshida and Toru Nagasawa

Microbial hydroxylation, which introduces regioselectively a hydroxyl group at a nonactivated carbon atom of alicyclic compounds, is an attractive and promising method for the synthesis of useful fine chemicals. Recently, we found that Streptomyces griseoplanus AC122 catalyzed highly regioselective hydroxylation of adamantane.1 Through hydroxylation of adamantane by S. griseoplanus AC122 cells, 1-adamantanol was synthesized in the culture broth (Figure 12.5). 12.4.1 12.4.1.1 • • • • • • • • • • •

Procedure 1: Cultivation of S. griseoplanus AC122 Materials and Equipment

Glucose (0.4 g) malt extract (1 g) yeast extract (0.4 g) tap water 100 mL malt extract (1 g) yeast extract (1 g) magnesium sulfate heptahydrate (0.012 g) tap water 100 mL one 50 mL test tube with a poromeric silicone plug one shaking-flask with a cotton plug, 500 mL reciprocal shaker.

12.4.1.2

Procedure

1. Glucose (0.4 g), malt extract (1 g) and yeast extract (0.4 g) were dissolved with water and the volume was adjusted to 100 mL with tap water. The seed culture medium (4 mL) was placed in a 50 mL test tube. Seed culture of S. griseoplanus AC122 was carried out for 48 h at 28 C with reciprocal shaking (115 strokes per minute). 2. Malt extract (1 g), yeast extract (1 g) and magnesium sulfate heptahydrate (0.012 g) were dissolved with water and the volume was adjusted to 100 mL with tap water. The culture medium (30 mL) was placed in a 500 mL shaking-flask with a cotton plug and sterilized (121 C, 20 min). The seed culture broth was transferred to a 500 mL shakingflask containing 30 mL of the culture medium. Cultivation was carried out for 48 h at 28 C and 115 strokes per minute. OH S. griseoplanus AC122

32% yield

Figure 12.5

Hydroxylation of adamantane using Streptomyces cells

368


12.4.2

Procedure 2: Synthesis of 1-Adamantanol OH

12.4.2.1 • • • • • • • • •


Adamantane (41 mg, 0.3 mmol) Tween 60 (900 mg) ethyl acetate anhydrous magnesium sulfate n-hexane and ethyl acetate filter paper silica gel (Wakogel C300 45–75 mm), 15 g one 300 mL separatory funnel rotary evaporator.

12.4.2.1

Procedure

1. Adamantane (40.9 mg, 0.3 mmol) and Tween 60 (900 mg) were added to 30 mL of culture medium after 48 h of cultivation. The conversion of adamantane was carried out for 72 h with reciprocal shaking (115 strokes per minute) at 28 C. 2. The cells were removed from the culture broth by centrifugation (12 000 rpm, 30 min) and the supernatant was extracted with ethyl acetate. The organic layer was collected, dried over anhydrous magnesium sulfate and concentrated using a rotary evaporator. The crude product was purified by a silica-gel column chromatography using eluents (the mixture of n-hexane and ethyl acetate was used stepwise at a ratio (v/v) of 1:0, 8:1, 4:1 and 2:1) to give 1-adamantanol (13 mg, 32 % yield). 1

H NMR (500 MHz, CDCl3) 1.47 (1H, s), 1.62 (6H, dd, J 24.3, 12.3 Hz) 1.71 (6H, d, J 2.3 Hz), 2.14 (3H, s); 13C NMR (125 MHz, CDCl3) 30.7, 36.0, 45.3, 68.2; electron impact mass spectrometry m/z (%) 152 (Mþ, 29.4), 95 (58.9). 12.4.3

Conclusion

The procedure is an approach for the synthesis of 1-adamantanol from adamantane by a green bioprocess.

Reference and Note 1. Mitsukura, K., Kondo, Y. Yoshida T. and Nagasawa, T., Regioselective hydroxylation of adamantane by Streptomyces griseoplanus cells. Appl. Microbiol. Biotechnol., 2006, 71, 502. Adamantane-hydroxylating strains such as Dothiora phaeosperma and Streptomyces griseus IFO3237 can be utilized for 1-adamantanol production.

12.5 Enantioselective Benzylic Microbial Hydroxylation of Indan and Tetralin

12.5

369

Enantioselective Benzylic Microbial Hydroxylation of Indan and Tetralin Renata P. Limberger, Cleber V. Ursini, Paulo J.S. Moran and J. Augusto R. Rodrigues

Benzylic microbial hydroxylations of hydrocarbons have been found to be an important tool in organic chemistry. Their efficiency, specificity and environmentally benign conditions make this approach superior to many chemical-based methods,1 especially in view of unsolved chemical problems, such as a certain lack of control and predictability of the product structures and the expense of oxidizing reagents.2 Recently, we have described a screening of 15 strains of bacteria and fungi targeted at the production of specific hydroxylated benzylic derivatives of indan and tetralin.3 Among the cultures screened, Mortierella isabellina CCT3498, Mortierella ramanniana CCT4428 and Beauveria bassiana CCT3161 (from ATCC 7159) were shown to mediate the respective conversions of the hydrocarbons into 1-indanol and 1-tetralol. The most satisfactory results were achieved with M. isabellina, which afforded (R)-1-indanol (78 % conversion, 64 % yield, 86 % ee) after a 2-day incubation and (R)-1-tetralol (50 % conversion, 38 % yield, 92 % ee) in a 4-day incubation. Overoxidation of 1-indanol and 1-tetralol during the reactions resulted in the formation of 1-indanone and 1-tetralone respectively. 12.5.1

Procedure 1: Synthesis of (R)-1-Indanol OH

O

M. isabellina

indian

12.5.1.1

(R)-1-indanol 64% yield

1-indanone


• Indan (95 %, Acros) (100 mg, 0.846 mmol, 2.0 mL of ethanolic solution at 50 mg mL1) • M. isabellina CCT3498 strain (Tropical Culture Collection, Andre Tosello Research Foundation)4 • potato–dextrose–carrot broth (PDCB, 200 mL)5 • pH 6.0 potassium phosphate buffer (18.4 g KH2PO4 and 4.025 g Na2HPO4 in 1 L) (200 mL) • ethyl acetate (500 mL, Merck) • hexane (700 mL, Merck) • sodium chloride, saturated aqueous solution • silica gel (400–200 mesh, Aldrich) (25 cm height in glass column) • anhydrous sodium sulfate (5 g) • two 500 mL Erlenmeyer flasks

370


• • • • • • • • • • • • •

orbital shaker centrifuge 500 mL filter flask Bu¨chner funnel filter paper magnetic stirring plate 250 mL separating funnel rotary evaporator equipment for continuous liquid–liquid extraction equipment for flash column chromatography using a glass column (2.5 cm 25 cm) thin-layer chromatography (TLC) plates (silica gel 60 F254, Merck) gas chromatograph–mass spectrometer (GC–MS) J&W Scientific HP-5 (30 m 0.25 mm 0.25 mm) or Supelco Simplicity 1 (30 m 0.25 mm 0.25 mm) fused-silica capillary column • Marcherey 212117/91 Hydrodex- 3P (25 m 0.25 mm 0.25 mm) fused silica capillary column • polarimeter • nuclear magnetic resonance spectrometer. 12.5.1.2

Procedure

1. A culture of M. isabellina CCT3498 was aseptically transferred into conical Erlenmeyer flasks (500 mL) containing 200 mL of sterile PDCB5 and kept on a rotary shaker (150 rpm) at 30 C for 3 days to acquire biomass. 2. Microbial biomass was harvested by centrifugation (5000 rpm, 10 min) and 30 g (wet weight) was transferred to clean 500 mL conical flasks containing 200 mL of 0.2 ionic strength potassium phosphate buffer solution at pH 6.0. Attention: to avoid the lack of efficiency, new cultures of M. isabellina should be used and a careful control of incubation conditions (temperature, pH and medium) is necessary. 3. 100 mg of indan (2.0 mL of ethanolic solution at 50 mg mL1) was added. 4. The flask was returned to the shaker (150 rpm) at 30 C for 2 days. 5. After 2 days, three 5.0 mL portions of the incubation mixture were harvested by vigorous shaking and extraction with 5.0 mL of ethyl acetate. If necessary, a centrifugation procedure (3 min at 3000 rpm) was used to break the emulsion. 6. The organic fraction was collected and 1 mL of the solution was submitted to the GC–MS for qualitative and chiral analysis to certify conversion to 1-indanol. 7. For qualitative analyses, the GC system was equipped with a J&W Scientific HP-5 or a Supelco Simplicity 1 fused-silica capillary column. Injector and detector temperatures were set at 220 C and 240 C respectively; the oven temperature was programmed from 60 to 230 C at 40 C min1. Helium was employed as carrier gas (1 mL min1). Compound identification was based on a comparison of mass spectra with those of synthetic racemic and enantiomeric-enriched samples. The retention times for indan, 1-indanol and 1-indanone were 4.7 min, 5.9 min and 6.2 min respectively. 8. For chiral analyses the GC system was equipped with a Marcherey 212117/91 Hydrodex- 3P fused-silica capillary column. The oven temperature was


9.

10. 11. 12.

13.

371

programmed from 100 to 210 C at 10 C min1. Injector and detector temperatures were set at 200 C and 240 C respectively. Hydrogen was employed as carrier gas (1 mL min1). Under these conditions, the retention times obtained for (S)-1-indanol and (R)-1-indanol were 8.07 min and 8.11 min respectively. After the incubation time, the culture was harvested and filtered. The filtrate was saturated with a sodium chloride saturated aqueous solution, stirred for 3–4 h at room temperature and then extracted six times with ethyl acetate (20 mL). The organic phase was saved and the sodium chloride saturated aqueous solution remaining was extracted again by a continuous liquid–liquid process at 50 C for 24 h. The combined organic extracts were dried over anhydrous sodium sulfate and concentrated in vacuum after filtration. The crude residue was purified by flash column chromatography on silica gel using 300 mL portions of hexane:ethyl acetate (90:10 and 80:20); 10 mL fractions were collected, giving indanone with a 9:1 ratio and 1-indanol (64 mg, 0.541 mmol) with 8:2 ratio. The isolated 1-indanol was collected and 1 mL of the solution was submitted for GC–MS analysis, as described above, and the compound identity was confirmed by nuclear magnetic resonance spectrometry.7,8

(R)-1-Indanol was isolated as a white solid in 64 % yield. M.p. 67–68.0 C; 9,10 m.p. 72 C, []22 []23 D ¼ 25 (c ¼ 0.41, CHCl3), 88 % ee. Lit.: D ¼ þ34 (c ¼ 1.895, 1 CHCl3) for (S) enantiomer. H NMR (CDCl3, 300 MHz): 7.46–7.42 (m, 1H, Ph), 7.24–7.20 (m, 2H, Ph), 7.14–7.10 (m, 1H, Ph), 5.27 (br t, J ¼ 5.9 Hz, 1H, CHOH), 3.07 (ddd, J ¼ 16.2 Hz, J ¼ 8.4 Hz, J ¼ 4.8 Hz, 1H, CHHCH2CH(OH)), 2.84 (br dd, J ¼ 16.2 Hz, J ¼ 7.0 Hz, 1H, CHHCH2C(OH)H), 2.51 (m, 1H, CHHC(OH)H), 1.97 (m, 1H, CHHC(OH)H), 1.75 (br s, 1H, OH). MS (electron impact (EI)): m/z (relative intensity) 134 (Mþ, 51), 133 (100), 117 (12), 116 (14), 115 (28), 105 (30), 103 (12), 91 (32), 89 (9), 79 (25), 78 (15), 77 (45), 74 (2), 66 (16), 65 (22), 63 (25), 57 (25), 55 (32), 53 (9), 52 (13), 51 (57), 50 (29). Racemic standards of 1-indanol, to be used for chiral GC analysis, can be prepared by treatment of indanone with NaBH4 in MeOH, as described by Aina et al.6 Enantiomerically enriched samples of 1-indanol, used to determine the enantiomer specificity by GC, can be prepared by hydrogen transfer ruthenium-catalysed reduction, as described by Ursini et al.7 12.5.2

Procedure 2: Synthesis of (R)-1-Tetralol OH

tetralin

(R)-1-tetralol 38% yield 92% ee

O

1-tetralone

372


12.5.2.1


• -Tetralin (99%, Sigma–Aldrich) (100 mg, 0.756 mmol, 2.0 mL of ethanolic solution at 50 mg mL1) • M. isabellina CCT3498 strain (Tropical Culture Collection, Andre Tosello Research Foundation)4 • PDCB (200 mL)5 • two 500 mL Erlenmeyer flasks • pH 7.0 potassium phosphate buffer (4.71 g KH2PO4 and 7.85 g Na2HPO4 in 1 L) (200 mL) • ethyl acetate (500 mL, Merck) • hexane (700 mL, Merck) • sodium chloride, saturated aqueous solution • anhydrous sodium sulfate (5 g) • silica gel (400–200 mesh, Aldrich) (23 cm height in glass column) • orbital shaker • centrifuge • 500 mL Kitasato • Bu¨chner funnel • filter paper • magnetic stirring plate • 250 mL Separating funnel • rotary evaporator • equipment for continuous liquid–liquid extraction • equipment for flash column chromatography using a glass column (2.5 cm 25 cm) • TLC plates (silica gel 60 F254, Merck) • GC–MS • J&W Scientific HP-5 (30 m 0.25 mm 0.25 mm) or Supelco Simplicity 1 (30 m 0.25 mm 0.25 mm) fused-silica capillary column • Marcherey 212117/91 Hydrodex- 3P (25 m 0.25 mm 0.25 mm) fused-silica capillary column • polarimeter • nuclear magnetic resonance spectrometer. 12.5.2.2

Procedure

1. A culture of M. isabellina CCT3498 was aseptically transferred to conical Erlenmeyer flasks (500 mL) containing 200 mL of sterile PDCB5 and kept on a rotary shaker (150 rpm) at 30 C for 3 days to acquire biomass. 2. Microbial biomass was harvested by centrifugation (5000 rpm, 10 min) and 30 g (wet weight) was transferred to clean 500 mL conical flasks containing 200 mL of 0.2 ionic strength potassium phosphate buffer solutions at pH 7.0. Attention: to avoid the lack of efficiency, new cultures of M. isabellina should be used and a careful control of incubation conditions (temperature, pH and medium) is necessary. 3. 100 mg of tetralin (2.0 mL of ethanolic solution at 50 mg mL1) was added. 4. The flask was returned to the shaker (150 rpm) at 30 C for 4 days.


373

5. After 4 days, three 5.0 mL portions of the incubation mixture were harvested and submitted to vigorous shaking and extraction with 5.0 mL of ethyl acetate. If necessary, a centrifugation procedure (3 min at 3000 rpm) was used to break the emulsion. 6. The organic fraction was collected and 1 mL of the solution was submitted to the GC–MS for qualitative and chiral analysis to certify the conversion to 1-tetralol. 7. For qualitative analyses, the GC system was equipped with a J&W Scientific HP-5 or a Supelco Simplicity 1 fused-silica capillary column. Injector and detector temperatures were set at 220 C and 240 C respectively; the oven temperature was programmed from 60 to 230 C at 40 C min1. Helium was employed as carrier gas (1 mL min1). Compound identification was based on a comparison of mass spectra with those of synthetic racemic and enantiomeric-enriched samples. The retention times for tetralin, 1-tetralol and 1-tetralone were 5.6 min, 6.5 min and 6.6 min respectively. 8. For chiral analyses the GC system was equipped with a Marcherey 212117/91 Hydrodex- 3P fused-silica capillary column. The oven temperature was programmed from 100 to 210 C at 10 C min1. Injector and detector temperatures were set at 200 C and 240 C respectively. Hydrogen was employed as carrier gas (1 mL min1). Under these conditions, the retention times obtained for (S)-1-tetralol and (R)-1-tetralol were 9.6 min and 9.7 min respectively. 9. After the incubation time, the culture was harvested and filtered. The filtrate was saturated with a sodium chloride saturated aqueous solution, stirred for 3–4 h at room temperature and extracted six times with ethyl acetate (20 mL). 10. The organic phase was saved and the sodium chloride saturated aqueous solution remaining was extracted again by continuous liquid–liquid process at 50 C for 24 h. 11. The combined organic extracts were dried over anhydrous sodium sulfate and concentrated in vacuum after filtration. 12. The crude residue was purified by flash column chromatography on silica gel using 300 mL portions of hexane:ethyl acetate (90:10 and 80:20); 10 mL fractions were collected, giving 1-tetralone with 9:1 ratio and 1-tetralol (38 mg, 0.287 mmol) with 8:2 ratio. 13. The isolated 1-tetralol was collected and 1 mL of the solution was submitted for GC– MS analysis, as described above; compound identity was confirmed by nuclear magnetic resonance spectrometry.7,8 (R)-1-Tetralol was isolated as a colourless oil in 38 % yield. []22 D ¼ 34.0 ¼ þ34.4 (c ¼ 1.01, CHCl ) for S enan(c ¼ 2.12, CHCl3), 92 % ee. Lit.:11,12 []25 3 D tiomer. 1H NMR (CDCl3, 300 MHz): 7.46–7.42 (m, 1H, Ph), 7.24–7.20 (m, 2H, Ph), 7.14–7.10 (m, 1H, Ph), 4.79 (apparent t, J ¼ 4.4 Hz, 1H, CHOH), 2.85–2.65 (m, 2H, CH2), 2.05–1.75 (m, 5H, CH2, CH2, OH). MS (EI): m/z (relative intensity) 148 (Mþ, 18), 147 (25), 131 (18), 129 (43), 128 (20), 127 (13), 121 (8), 120 (80), 119 (67), 115 (28), 105 (47), 104 (15), 92 (20), 91 (100), 90 (15), 89 (15), 79 (10), 78 (30), 77 (34), 66 (10), 65 (47), 64 (30), 63 (41), 62 (10), 60 (10), 57 (11), 55 (10), 53 (14), 52 (16), 51 (69), 50 (28), 43 (23), 41 (31), 40 (12). Racemic standards of 1-tetralol, to be used for chiral GC analysis, can be prepared by treatment of tetralone with NaBH4 in MeOH, as described by Aina et al.6

374


Enantiomerically enriched samples of 1-tetralol, used to determine the enantiomer specificity by GC, can be prepared by hydrogen transfer ruthenium-catalysed reduction, as described by Ursini et al.7 12.5.3

Conclusion

The enantioselective benzylic hydroxylation of indan and tetralin can be achieved with M. isabellina, affording 78 % conversion to 1-indanol (64 % yield, 86 % (1R)- ee) in a 2day incubation and 52 % conversion to 1-tetralol (38 % yield, 92 % (1R)- ee) in a 4-day incubation. The good yields and ee allow their use in future scaling-up processes; however, to avoid the lack of efficiency, careful control of the temperature, pH and medium is necessary, since the reactions are strongly dependent on the incubation and reaction conditions. Tables 12.2 and 12.3 give details of some of the different incubation conditions/results and time-course analysis found in the benzylic hydroxylation of indan and tetralin mediated by M. isabellina CCT3498.

Table 12.2 Benzylic hydroxylation of indan and tetralin mediated by M. isabellina CCT3498 a Entry

Parameter

1 2 3 4 5

pH Incubation time (days) Relative conversion to alcohol (%)b Alcohol yield (%)c Ee (% R)d

Indan to 1-indanol

Tetralin to 1-tetralol

6.0 2 78 64 86

7.0 4 50 38 92

a

The microorganism was grown in PDCB at 30 C. The reactions were performed in buffer solutions, also at 30 C. Determined by GC on an HP-5 or a Supelco Simplicity 1 fused-silica capillary column. The percentage compositions were obtained from electronic integration measurements, without taking into account relative response factors. c The yields quoted are those of isolated, purified material. d Determined by chiral GC on Marcherey 212117/91 Hydrodex- 3P fused-silica capillary column. b

Table 12.3 Time-course analysis obtained in the benzylic hydroxylation of indan and tetralin (30 mg) by M. isabellina (3 g fresh weight) a Time

Day 1 Day 2 Day 3 Day 4 Day 5

Conversion (%) of substrate and products Indan

1-Indanol

1-Indanone

Tetralin

1-Tetralol

1-Tetralone

75 18 18 18 13

24 78 64 59 36

1 3 18 23 51

89 65 22 14 0

10 28 52 50 40

1 7 28 36 60

a The experiments were carried out in triplicate and analysed by GC on an HP-5 or a Supelco Simplicity 1 fused-silica capillary column. The percentage compositions were obtained from electronic integration measurements, without taking into account relative response factors.


375

References and Notes 1. Van Berkel, W.J.H., Kamerbeek, N.M. and Fraaije, M.W., Flavoprotein monooxygenases, a diverse class of oxidative biocatalysts. J. Biotechnol., 2006, 124, 670. 2. Burton, S.G., Oxidizing enzymes as biocatalysts. Trends Biotechnol., 2003, 21, 543. 3. Limberger, R.P., Ursini, C.V., Moran, P.J.S. and Rodrigues, J.A.R., Enantioselective benzylic microbial hydroxylation of indan and tetralin. J. Mol. Catal. B: Enzym. 2007, 46, 37. 4. Fundac˛aõ Andre´ Tosello de Pesquisa e Tecnologia, Rua Latino Celho 1301, 13087-1001 Campinas-SP, Brazil, http://www.fat.org.br. 5. PDCB was prepared by suspending cut-up unpeeled potatoes (100 g L1) and carrots (10 g L1) in purified water and heated to boiling in a microwave oven for 5–10 min. Dextrose (D-(þ)glucose) was added (30 g L1). The medium was sterilized by autoclaving for 20 min at 121 C and then decanting off the broth. The broth is clear to slightly opalescent and yellowish in colour. No pH adjustment was made. 6. Aina, G., Nasini, G. and Pava, O.V., Asymmetric bioreduction of racemic 5,6,7,8-tetrahydro-8methyl-1,3-dimethoxynaphthalen-6-one to the corresponding chiral -tetralols. J. Mol. Catal. B: Enzym., 2001, 11, 367. 7. Ursini, C.V., Dias, G.H.M. and Rodrigues, J.A.R., Ruthenium-catalyzed reduction of racemic tricarbonyl(6-aryl ketone)chromium complexes using transfer hydrogenation: a simple alternative to the resolution of planar chiral organometallics. J. Organomet. Chem., 2005, 690, 3176. 8. Boyd, D.R., Sharma, N.D., Boyle, R., Evans, T.A., Malone, J.F., McCombe, K.M., Dalton, H. and Chima, J., Chemical and enzyme-catalysed syntheses of enantiopure epoxide and diol derivatives of chromene, 2,2-dimethylchromene, and 7-methoxy-2,2-dimethylchromene (precocene-1). J. Chem. Soc. Perkin Trans. 1, 1996, 1757. 9. Brand, J.M., Cruden, D.L., Zylstra, G.J. and Gibson, D.T., Stereospecific hydroxylation of indan by Escherichia coli containing the cloned toluene dioxygenase genes from Pseudomonas putida F1. Appl. Environ. Microbiol., 1992, 58, 3407. 10. Jaouen, G. and Meyer, A., Facile syntheses of optically active 2-substituted indanones, indanols, tetralones, and tetralols via their chromium tricarbonyl complexes. J. Am. Chem. Soc., 1975, 97, 4667. 11. Boyd, D.R., McMordie, R.A S., Sharma, N.D., Dalton, H., Williams, P. and Jenkins, R.O., Stereospecific benzylic hydroxylation of bicyclic alkenes by Pseudomonas putida: isolation of (þ)-R-1-hydroxy-1,2-dihydronaphthalene, an arene hydrate of naphthalene from metabolism of 1,2-dihydronaphthalene. J. Chem. Soc. Chem. Commun., 1989, 339. 12. Palmer, M.J., Kenny, J.A., Walsgrove, T., Kawamoto, A.M. and Wills, M., Asymmetric transfer hydrogenation of ketones using amino alcohol and monotosylated diamine derivatives of indane. J. Chem. Soc. Perkin Trans. 1, 2002, 416.

376

12.6


Stereospecific Biotransformation of (R,S)-Linalool by Corynespora cassiicola DSM 62475 into Linalool Oxides Marco Antonio Mirata and Jens Schrader

The biotransformation of (R,S)-linalool by fungi is a useful method for the preparation of natural linalool oxides.1 The stereospecific conversion of (R,S)-linalool by Corynespora cassiicola DSM 62475 led to 5R-configured furanoid linalool oxides and 5S-configured pyranoid linalool oxides, both via 6S-configured epoxylinalool as postulated intermediate (Figure 12.6). The biotransformation protocol affords an almost total conversion of the substrate with high enantioselectivities and a molar conversion yield close to 100 % (Table 12.4). Pure linalool oxides are of interest for lavender notes in perfumery.1

5 2 O

HO HO

HO

furanoid cis-(2S,5R ) linalool oxide

C. cassiicola DSM 62475 O

(3S )-linalool

HO 5

2 O

(3S, 6S )-epoxylinalool

pyranoid cis-(2S, 5S ) linalool oxide

5 2 O

HO HO

HO

furanoid trans-(2R,5R ) linalool oxide

C. cassiicola DSM 62475

HO O

5 2

(3R )-linalool

(3R, 6S )-epoxylinalool

O

pyranoid trans-(2R,5S ) linalool oxide

Figure 12.6 Stereospecific biotransformation of (R,S)-linalool by C. cassiicola via the (6S)-configured epoxylinalools as postulated intermediates

12.6.1 12.6.1.1 • • • •

Procedure 1: Preparation of Spores Suspension of C. cassiicola DSM 62475 Materials and Equipment

Malt extract (30 g) glucose (10 g) peptone (4 g) agar (17 g)

12.6 Stereospecific Biotransformation of (R,S)-Linalool

• • • • • • • • • •

377

yeast extract (3 g) NaCl (8.5 g) Tween 80 (1 g) stock culture of C. cassiicola DSM 62475 distilled water 2 L acetic acid (AcOH, >99.8 %) one Petri dish one Falcon tube 50 mL incubator Drigalski spatula.

12.6.1.2

Procedure

1. C. cassiicola DSM 62475 was taken from stock culture and incubated for 15 days at 25 C on malt extract agar plate (consisting of 30 g malt extract, 3 g peptone and 17 g agar in 1 L distilled water adjusted to pH 5.6 with AcOH and sterilized at 121 C for 20 min). 2. A spore suspension was prepared by resuspending the spores from the agar plate in 15 mL physiological aqueous solution (8.5 g NaCl, 1 g peptone, and 1 g Tween 80 in 1 L distilled water, sterilized at 121 C for 20 min) with a Drigalski spatula. The spore suspension was diluted to a concentration of approximately 2.5 107 spores/mL and stored in a 50 mL Falcon tube at 4 C. 12.6.2 12.6.2.1 • • • • • • • • • • • •

Procedure 2: Fed-batch Biotransformation of (R,S)-Linalool by C. cassiicola DSM 62475 Materials and Equipment

Malt extract (30 g) glucose (10 g) peptone (10 g) yeast extract (30 g) spore suspension (R,S)-linalool stock solution (3 % w/v in ethanol 99 %) glucose aqueous solution 1.1 kg L1 acetic acid (AcOH, >99.8 %) one Erlenmeyer flask 300 mL one Erlenmeyer flask 2 L one bottle 1 L shaking incubator.

12.6.2.2

Procedure

1. Malt extract (30 g), glucose (10 g), peptone (10 g) and yeast extract (30 g) were dissolved with water and the volume was adjusted to 1.0 L with distilled water; the pH was adjusted to 6.4 with AcOH. The resulting solution (MYB medium) was sterilized (121 C, 20 min) and stored at 4 C.

378


Table 12.4 Fed-batch biotransformation of (R,S)-linalool by C. cassiicola DSM 62475 using Procedures 1 and 2. Of 340 mg L1 (R,S)-linalool added, 96 % was consumed Products from (R, S)-linalool Furanoid trans-(2R,5R)-linalool oxide Furanoid cis-(2S,5R)-linalool oxide Pyranoid trans-(2R,5S)-linalool oxide Pyranoid cis-(2S,5S)-linalool oxide

Molecular yield (%)

Concentration (mg/L)

Ee (%)

43.0 41.7 4.7 9.2

153 148 17 33

>95 >99 >80 >97

2. For the preparation of the preculture, 50 mL of MYB medium was placed in a sterile 300 mL Erlenmeyer flask and inoculated with 500 mL spore supension of C. cassiicola DSM 62475. The Erlenmeyer flask was incubated and shaken for 24 h at 25 C and 130 rpm. 3. A sterile 2 L Erlenmeyer flask was filled with 450 mL MYB medium containing 2.5 mL of (R,S)-linalool stock solution (150 mg L1 linalool) and inoculated with the resulting preculture (step 2) of C. cassiicola DSM 62475. The Erlenmeyer flask was incubated and shaken for 72 h at 25 C and 130 rpm. A volume of 1.7 mL (R,S)-linalool stock solution (100 mg L1 linalool) and 2 mL glucose aqueous solution (5 g L1) were fed after 24 and 48 h cultivation. 12.6.3

Conclusion

The procedure is very easy to reproduce and to scale up. Bioconversion products can be easily isolated by evaporation of the extraction solvent (e.g. tert-butyl methyl ether). Table 12.4 summarizes the product concentrations, molecular conversion yields and enantioselectivities obtained during linalool biotransformation with C. cassiicola DSM 62475.

Reference 1. Schrader, J. and Berger, R.G., Biotechnological production of terpenoid flavor and fragrance compounds. In Biotechnology, vol. 10, 2nd edn, Rehm, H.-J. and Reed, G. (eds). Wiley-VCH: Weinheim, 2001, pp. 377–383.

12.7 The Biocatalytic Synthesis of 4-Fluorocatechol from Fluorobenzene

12.7

379

The Biocatalytic Synthesis of 4-Fluorocatechol from Fluorobenzene Louise C. Nolan and Kevin E. O’Connor*

The microbial synthesis of organic compounds is a useful method for the preparation of valuable compounds such as substituted catechols. Here, we describe two approaches to the biological formation of 4-fluorocatechol from fluorobenzene. First, we describe the biotransformation of fluorobenzene to 4-fluorocatechol using whole cells of Pseudomonas mendocina KR1 expressing toluene-4-monooxygenase (T4MO). Second, we use whole cells expressing T4MO in tandem with the enzyme tyrosinase sourced commercially from mushrooms to further improve catechol formation. F

F T4MO

F

F Ascorbic acid

T4MO Tyrosinase OH OH

Fluorobenzene

12.7.1 12.7.1.1 • • • • • • • • • • • • • • • • • • • • • •

4-Fluorophenol

OH 4-Fluorocatechol

O O 4-Fluoroquinone

Procedure 1: Growth Medium and Buffers Materials and Equipment

Sodium ammonium phosphate tetrahydrate (NaNH4HPO44H2O (35.0 g)) potassium phosphate dibasic trihydrate (K2HPO43H2O (75.0 g)) potassium phosphate monobasic (KH2PO4 (37.0 g)) magnesium sulfate heptahydrate (MgSO47H2O (4.93 g)) ferrous sulfate heptahydrate (FeSO47H2O (2.78 g)) manganese chloride tetrahydrate (MnCl24H2O (1.98 g)) cobalt(II) sulfate heptahydrate (CoSO47H2O (2.81 g)) calcium chloride dihydrate (CaCl22H2O (1.47 g)) copper(II) chloride dehydrate (CuCl22H2O (0.17 g)) zinc sulfate heptahydrate (ZnSO47H2O (0.29 g)) hydrochloric acid (HCl) (37 %) biotin (20 mg) folic acid (20 mg) pyrodoxine hydrochloride (100 mg) riboflavin (50 mg) thiamine hydrochloride (50 mg) nicotinic acid (50 mg) pantothenic acid (50 mg) vitamin B12 (1 mg) 4-aminobenzoic acid (50 mg) DL-6,8-thioctic acid (50 mg) K2HPO4.3H2O (11.423 g)

380


• KH2PO4 (6.805 g) • glycerol • deionized water. 12.7.1.2

Procedure

1. Stock solution 1. 50 stock solution of E2 mineral salts medium was prepared as follows: NaNH4HPO44H2O (35.0 g) was dissolved in 100 mL deionized water using magnetic stirring. K2HPO43H2O (75.0 g) and KH2PO4 (37.0 g) were added to the solution and the volume adjusted to 200 mL with deionized water. The pH was adjusted to 7.0. This solution was stored unautoclaved on the bench. 2. Stock solution 2. MgSO47H2O (4.93 g) was dissolved in 20 mL of deionized water. This solution was sterilized by autoclaving and stored as a 1 M stock solution on the bench. 3. 100 mL of 1 M HCl was prepared by adding 8.35 mL of 37 % HCl to 91.65 mL of deionized water. 4. Stock solution 3. 100 stock solution of trace elements was prepared by dissolving FeSO47H2O (2.78 g), MnCl24H2O (1.98 g), CoSO47H2O (2.81 g), CaCl22H2O (1.47 g), CuCl22H2O (0.17 g) and ZnSO47H2O (0.29 g) in 1 M HCl. The final volume was adjusted to 1.0 L. This stock solution was stored at 4 C. 5. Stock solution 4. 100 stock solution of vitamins was prepared by dissolving biotin (20 mg), folic acid (20 mg), pyrodoxine hydrochloride (100 mg), riboflavin (50 mg), thiamine hydrochloride (50 mg), nicotinic acid (50 mg), pantothenic acid (50 mg), vitamin B12 (1 mg), 4-aminobenzoic acid (50 mg) and thioctic acid (50 mg) in deionized water. The volume was adjusted to 1.0 L. The solution was filtered, sterilized and stored as 10 mL aliquots at 20 C. 6. Stock solution 5. 1 M stock solution of potassium phosphate buffer was prepared by dissolving K2HPO43H2O (11.423 g) and KH2PO4 (6.805 g) in deionized water to a final volume of 100 mL. The pH was adjusted to 7.0. This 1 M stock solution was diluted to the desired concentration of 50 mM with deionized water. Buffers were stored at 0–4 C. 7. Stock solution 6. 60 % glycerol was prepared by mixing 12 mL glycerol with 8 mL deionized water. This solution was autoclaved and stored on the bench. 12.7.2 12.7.2.1 • • • • • • • • • •

Procedure 2: Storage, Cultivation and Harvesting of P. mendocina KR1 Materials and Equipment

Stock solution 1 (9 mL) deionized water (439 mL) stock solution 2 (450 mL) stock solution 3 (450 mL) stock solution 4 (450 mL) toluene (800 mL) 1 250 mL centre column Erlenmeyer flask with cotton wool plug 2 2 L centre column Erlenmeyer flask with cotton wool plugs New Brunswich Scientific C25 incubator shaker (Classic Series) stock solution 6 (250 mL)


381

• 15–20 1.5 mL sterile polypropylene tubes • Unicam (UV–vis) Helios thermo-spectrophotometer • Du Pont RC5C-plus fixed-angle centrifuge 12.7.2.2

Procedure

1. Overnight starter cultures of P. mendocina KR1 were grown in a 250 mL glass centre column (fused to the base of the flask) Erlenmeyer flask. Each flask contained stock solution 1 (1 mL) and deionized water (49 mL). This solution was autoclaved and cooled to room temperature before adding 50 mL of stock solution 2, 50 mL stock solution 3 and 50 mL stock solution 4. Toluene (200 mL) was added to the glass centre column as the sole source of carbon and energy supplied in the vapour phase. The flask was then inoculated with 100 mL of a freezer stock of P. mendocina KR1. Cultures were grown on a shaker table incubator at 30 C for 18 h at 200 rpm. 2. 750 mL of the above culture was added to stock solution 6 (250 mL) in 1.5 ml sterile tubes. The cultures were mixed gently before storing them at 80 C. These culture tubes were used as stocks for future inoculations. 3. Batch cultivation of P. mendocina KR1 was carried out in 2 2 L centre column Erlenmeyer flasks. Each flask contained stock solution 1 (8 mL) and deionized water (390 mL). This solution was autoclaved and cooled before adding 400 mL stock solution 2, 400 mL stock solution 3 and 400 mL stock solution 4. Toluene (600 mL) was added to the centre column. Overnight starter cultures were used to inoculate (2 % v/v) the growth medium. Cultures were incubated at 30 C shaking at 200 rpm. 4. Cells were harvested at an optical density (OD) (540 nm) of 0.7–0.8. During the harvest process, cells were kept on ice where possible. Cells were centrifuged at 16 200g for 10 min. Cell pellets were washed with 800 mL of ice-cold stock solution 5 and centrifuged as above. Cell pellets were combined and concentrated by resuspending them in a final volume of 50 mL of 50 mM stock solution 5. The OD (540 nm) was adjusted to 5 (1.5 mg cells dry weight (CDW)/mL). 12.7.3 12.7.3.1

Procedure 3: Biotransformation of Fluorobenzene by P. mendocina KR1 Materials and Equipment

Fluorobenzene (4.68 mL) ascorbic acid (176.12 mg) deionized water 1 % (w/v) D-glucose (0.5 g) 1 250 mL Erlenmeyer flask with cotton wool plug magnetic stirrer plate and magnetic bar 1 M HCl Eppendorf centrifuge 5810 R nylon filters (0.2 mm) high performance liquid chromatography (HPLC) vials and caps Hewlett Packard HP1100 instrument equipped with an Agilent 1100 series diode array detector • C18 Hypersil ODS 5 m HPLC column (125 mm 3 mm).

• • • • • • • • • • •

382


12.7.3.2

Procedure

1. 50 mL of the washed cell suspension of P. mendocina KR1 (1.5 mg CDW/mL) was transferred to a 250 mL Erlenmeyer flask and the contents brought to room temperature. 2. Ascorbic acid (176.12 mg) was dissolved in 1 mL of deionized water (1 M stock concentration). 3. 0.5 mL of 1 M ascorbic acid (10 mM final concentration) and D-glucose (0.5 g) were added to the cell suspension and the flask was placed on a magnetic stirrer. The contents were stirred magnetically at 200 rpm throughout the biotransformation. 4. 4.68 mL fluorobenzene (1 mM final concentration) was added to the flask and the flask was plugged with cotton wool. 5. The biotransformation was monitored by analysing samples taken periodically by HPLC. Samples (450 mL) were withdrawn from the biotransformation medium, acidified with 1 M HCL (50 ml) to stop the reaction and stored on ice for 30 min. All samples were centrifuged at 23 000g for 10 min at 4 C to remove the cell debris and the supernatant filtered into HPLC vials using nylon filters (0.2 mm). 6. Biotransformation samples were analysed by HPLC using a C18 Hypersil ODS 5m column (125 mm 3 mm) and a Hewlett Packard HP1100 instrument equipped with an Agilent 1100 series diode array detector. The samples were isocratically eluted using an aqueous phosphoric acid (0.1 % v/v)/methanol mix (70:30 (v/v)) at a flow rate of 0.5 mL min1. 7. After 120 min, whole cells of P. mendocina KR1 expressing T4MO activity transformed 1 mM fluorobenzene to 0.8 mM 4-fluorocatechol as a single product via 4-fluorophenol. 12.7.4

12.7.4.1 • • • • • • • •

Procedure 4: Biotransformation of Fluorobenzene by Whole Cells of P. mendocina KR1 Expressing T4MO in Tandem with a Cell-free Preparation of Tyrosinase from Mushroom Materials and Equipment

Fluorobenzene (23.4 mL) 1 % (w/v) D-glucose (0.5 g) 1 250 mL Erlenmeyer flask with cotton wool plug magnetic stirrer plate and magnetic bar Du Pont RC5C-plus fixed-angle centrifuge heated (30 C) oxygen electrode chamber mushroom (commercial) tyrosinase (10 mg) ascorbic acid (1 M stock).

12.7.4.2

Procedure

1. In a 250 mL Erlenmeyer flask, 50 mL of the washed cell suspension of P. mendocina KR1 (1.5 mg CDW/mL) and 0.5 g D-glucose were added and the contents brought to room temperature. 2. 23.4 mL of fluorobenzene (5 mM final concentration) was added to the flask and the flask was plugged with cotton wool. The contents were magnetically stirred at 200 rpm throughout the biotransformation.


383

3. After 105 min, the biotransformation contents were transferred to a centrifuge bucket and the cells spun out at 16 200g for 10 min at 4 C. 15 mL of the supernatant was transferred to an oxygen electrode chamber and brought to 30 C. Then, 15 mL 1 M ascorbic acid (1 mM final concentration) was added to the supernatant. 4. 10 mg mushroom tyrosinase was dissolved in stock solution 5 (1 mL) and kept on ice. 45 ml of mushroom tyrosinase (0.03 mg mL1 final concentration) was added to the supernatant to start the tyrosinase reaction and 0.03 mg mL1 was added every 15 min thereafter. In addition, 1 mM ascorbic acid was added every 5 min or until a colour change was observed. The reaction was stirred magnetically throughout. 5. The biotransformation was monitored by analysing samples by HPLC using the same sample preparation and HPLC analysis methods as described above (Procedure 3, Section 12.7.3). 6. After 120 min, tyrosinase transformed 1.8 mM 4-fluorophenol (produced by whole cells of P. mendocina KR1 expressing T4MO) to 1.3 mM 4-fluorocatechol. 12.7.5

Conclusion

The biotransformation of low levels of fluorobenzene (1 mM final concentration) to 4-fluorocatechol by whole cells of P. mendocina KR1 (1.5 mg CDW/mL) is easy to reproduce. Under these conditions, 4-fluorocatechol is formed as a single product in the biotransformation after 120 min (Table 12.5). Biotransformations with P. mendocina KR1 (1.5 mg CDW/mL) and higher concentrations of fluorobenzene (5 mM final concentration) result in the formation of 4-fluorophenol (1.8 mM) as a major product. In addition, minor products, namely 2-fluorophenol, 3-fluorophenol, 4-fluorocatechol and 3-fluorocatechol, are also formed. In the presence of ascorbic acid, tyrosinase has the ability to convert 4-fluorophenol (1.8 mM) to 4-fluorocatechol (1.3 mM). While this is a reproducible procedure, the 4-fluorocatechol does not accumulate as a single product (Table 12.5).

Table 12.5 Product formation in biotransformations with whole cells of P. mendocina KR1 expressing T4MO alone and in tandem with mushroom tyrosinase a Biotransformation conditions

2Fluorophenol

3Fluorophenol

4Fluorophenol

3Fluorocatechol

4Fluorocatechol

P. mendocina KR1 (1.5 mg CDW/mL) 1 mm fluorobenzene

ND

ND

ND

ND

0.8 mM

P. mendocina KR1 (1.5 mg CDW/mL), 5 mM fluorobenzene and mushroom tyrosinase in tandem

0.09 mM

0.08 mM

0.5 mM

0.03 mM

1.3 mM

a

ND: not detected.

384


References 1. Nolan, L.C. and O’Connor, K.E., Use of Pseudomonas mendocina, or recombinant Escherichia coli cells expressing toluene-4-monooxygenase, and a cell-free tyrosinase for the synthesis of 4-fluorocatechol from fluorobenzene. Biotechnol. Lett., 2007, 29, 1045. 2. Brooks, S.J., Doyle, E.M., Hewage, C., Malthouse, J.P.G. and O’Connor, K.E., Biotransformation of halophenols using crude cell extracts of Pseudomonas putida F6. Appl. Microbiol. Biotechnol., 2004, 64, 486.

12.8 Synthesis of Enantiopure (S)-Styrene Oxide by Selective Oxidation of Styrene

12.8

385

Synthesis of Enantiopure (S)-Styrene Oxide by Selective Oxidation of Styrene by Recombinant Escherichia coli JM101 (pSPZ10) Katja Buehler and Andreas Schmid

Selective oxidation of hydrocarbons is one of the most useful biotransformations for synthetic applications. Chemical counterparts often do not exist or lack the required regio- and enantioselectivity. In the respective reaction, recombinant Escherichia coli JM101 (pSPZ10) carrying and expressing the styAB genes encoding for the two-component styrene–monooxygenase from Pseudomonas sp. strain VLB120 is applied in a twoliquid-phase process for the highly enantioselective production of (S)-styrene oxide from toxic styrene (Figure 12.7).1,2 12.8.1 12.8.1.1

Procedure 1: Cultivation of the Seed Culture of Recombinant E. coli JM101 (pSPZ10) Materials and Equipment

• Luria–Bertani (LB) broth3 containing: – glucose 1 % (w/v) – kanamycin (50 mg L1) – deionized water – stored culture of E. coli JM101 (pSPZ10) • one 10 mL test tube with cap • sterile filters 0.2 mm pore size • shaker. 12.8.1.2

Procedure

1. The LB medium and the test tube were sterilized by autoclaving (121 C, 20 min), while the glucose and the kanamycin were dissolved separately in water and sterilized by filtration through a 0.2 mM filter. After allowing the LB medium to cool to room temperature, glucose and kanamycin solution were added in the appropriate amounts. 2. 5 mL of the thus-prepared medium were transferred to a sterile 10 mL test tube. The solution was inoculated with one colony of E. coli JM101 (pSPZ10) and left for incubation on a shaker at 250 rpm and 30 C overnight. O E. coli JM101 (pSPZ10)

ee > 99% Yield: > 76 %

Figure 12.7 Enantioselective epoxidation of styrene to (S)-styrene oxide utilizing recombinant E. coli JM101 (pSPZ10) as biocatalyst

386


12.8.2 12.8.2.1

Procedure 2: Cultivation of the Preculture of Recombinant E. coli JM101 (pSPZ10) Materials and Equipment

• M9* mineral salt medium: – disodium hydrogen phosphate dihydrate (25.5 g) – potassium dihydrogen phosphate (9.0 g) – ammonium chloride (1.0 g) – sodium chloride (0.5 g) – deionized water (1 L) – magnesium sulfate (240.5 mg) – kanamycin (50 mg) – thiamine (10 mg) – glucose (5 g) • US* trace element solution ( 1000): – hydrochloric acid (1 M) – manganese chloride tetrahydrate (1.5 g) – zinc sulfate (1.05 g) – boric acid (0.3 g) – sodium molybdate dihydrate (0.25 g) – copper(II) chloride dihydrate (0.15 g) – sodium ethylenediaminetetraacetic acid dihydrate (0.84 g) – calcium chloride dihydrate (4.12 g) – ferrous sulfate heptahydrate (4.87 g) • one 1000 mL shake flask with baffles • sterile filters 0.2 mm pore size • shaker. 12.8.2.2

Procedure

1. Disodium hydrogen phosphate dihydrate (25.5 g), potassium dihydrogen phosphate (9.0 g), ammonium chloride (1.0 g) and sodium chloride (0.5 g) were dissolved in water and the volume adjusted to 900 mL. The solution was sterilized by autoclaving (121 C, 20 min) and allowed to cool to room temperature. Magnesium sulfate (240.5 mg), kanamycin (50 mg), thiamine (10 mg) and glucose (5 g) were dissolved in water and adjusted to 100 mL volume. This mixture was sterilized by filtration through a 0.2 mm filter and added to the salt solution. 2. For the US* trace element solution ( 1000) all compounds were subsequently dissolved in 1 L of 1 M hydrochloric acid. The solution was sterilized by filtration through a 0.2 mm filter. 1 mL of this solution was added to 1 L of M9* medium prior to usage. 3. 100 mL of the ready-to-use M9* medium was transferred to a sterile 1000 mL shake flask with baffles. This solution was inoculated with 1 mL of freshly grown seed culture of E. coli JM101 (pSPZ10) (see Procedure 1, Section 12.8.1) and incubated overnight at 250 rpm and 30 C.


12.8.3 12.8.3.1 • • • • • • • • • • • • • • •

387

Procedure 3: Batch and Fed-batch Cultivation of Recombinant E. coli JM101 (pSPZ10)2 Materials and Equipment4

Dipotassium hydrogen phosphate trihydrate (15.9 g) potassium dihydrogen phosphate (4.0 g) disodium hydrogen phosphate dodecahydrate (7 g) ammonium sulfate (1.2 g) ammonium chloride (0.2 g) magnesium sulfate heptahydrate (1 g) yeast extract (5 g ) L-leucine (0.6 g) L-proline (0.6 g) deionized water (1 L) kanamycin (50 mg) thiamine (10 mg) glucose (5 g) US*trace element solution (1 mL) glucose feed medium:

– glucose (450 g L1) – magnesium sulfate heptahydrate (9 g L1) – dissolved in deionized water • bioreactor specifications: – lab-scale fermenter with a working capacity of 2.6 L made out of stainless steel, glass and Viton sealing – baffles and two six-bladed impellers allowing a stirrer speed of up to 3000 rpm – temperature control accomplished by using a Testoterm type II sensor and connecting the fermenter to a heating/cooling system – pH control connected to a rotary peristaltic pump to feed the titrants – in situ autoclavable amperometric probe (Pt/Ag) equipped with a fluoroethylene propylene (25 mm) membrane for dissolved oxygen tension (DOT) control – sterile filters (0.2 mm) for the air supply – thermostatted bubble column (i.d. 70 mm; h ¼ 350 mm) – foam probe – computer-controlled peristaltic pump and a microcomputer-connected balance to control the feed – LabTech Notebook software to control process parameters (pH, oxygen tension, air flow rate, glucose feed) – polypropylene glycol (20 % v/v), PP-G200 – ammonia (25 %) – phosphoric acid (25 %). 12.8.3.2

Procedure

1. Dipotassium hydrogen phosphate trihydrate (15.9 g), potassium dihydrogen phosphate (4.0g), disodium hydrogenphosphate dodecahydrate (7 g), ammonium sulfate

388

2.

3.

4.

5.


(1.2 g), ammonium chloride (0.2 g), yeast extract (5 g), L-leucine (0.6 g) and L-proline (0.6 g) were dissolved in water and the volume adjusted to 900 mL. The fermenter was assembled and filled with 900 mL of the medium prepared above (see step 1). This setup was then sterilized by autoclaving (121 C; 20 min) and allowed to cool to room temperature. Then, the fermenter was properly connected to air supply, pH titrants (ammonia solution and phosphoric acid) and anti-foam PP-G200, which was sterilized prior to usage. Magnesium sulfate (1 g), kanamycin (50 mg), thiamine (10 mg) and glucose (5 g) were dissolved in water and adjusted to 100 mL volume. This mixture was sterilized by filtration through a 0.2 mm filter and was added together with 1 mL of US* trace element solution (see Procedure 2, Section 12.8.2) to the fermenter. Batch growth was started by inoculating the fermenter with 100 mL of a freshly grown preculture of E. coli JM101 (pSPZ10) (see Procedure 2, Section 12.8.2). The pH was kept at 7.1, the aeration rate was set to 2 L min1 and the stirrer speed and temperature were 1300 rpm and 30 C respectively. The glucose feed was started as soon as the DOT increased significantly, indicating that the carbon source in the culture medium was consumed. The stirrer rate was increased to 2400 rpm. All other parameters were kept constant. Feed rate was maintained at 4.5 g glucose/Laq/h throughout the fed-batch.

12.8.4 12.8.4.1

Procedure 4: Biotransformation of Styrene into (S)-Styrene Oxide by recombinant E. coli JM101 (pSPZ10) Materials and Equipment

• Bis(2-ethylhexyl)phthalate (BEHP, 1 L) • n-octane (99 %) • styrene (99 %). 12.8.4.2

Procedure

1. The BEHP was supplemented with 1 % (v/v) octane for induction of the styAB genes and 4 % (v/v) styrene as substrate. 2. 1 h after the fed-batch was started (Procedure 3, Section 12.8.3) the organic phase was added to the bioreactor. The biotransformation was left running for 12 h, maintaining constant conditions as described in Procedure 3 (Section 12.8.3). 3. The organic phase was separated from the aqueous phase containing biomass by centrifugation. Epoxide products were recovered from the organic phase by vacuum distillation. 4. Ee was determined by gas chromatography (GC) on a Supelco Beta-DEX 120 column (fused-silica capillary column, 30 m, 0.25 mm inner diameter, 0.25 mm film thickness; Supelco, Buchs, Switzerland) with split injection (20:1) and an isothermal oven temperature profile at 90 C for separation of styrene oxide enantiomers. 12.8.5

Conclusion

During the overall biotransformation, the product formation rate reached a maximum of 61 U g1 cells dry weight (CDW) and decreased to 27 U g1 CDW towards the end of the process. This resulted in a final product concentration of 306 mM (S)-styrene oxide in the


389

organic phase. By applying the two-liquid phase concept, inhibition by substrate and product toxicity could be circumvented. Table 12.6 gives an overview of the different substrates, which are epoxidized with high enantiomeric excess by this biocatalyst. Table 12.6 Substrates and products with the corresponding yields and ee–values for the biocatalyst E. coli JM101 (pSPZ19) 5 Substratea

Productb

Yield (%)

Ee (%)

76.3

99.5

46.5

99.9

74.8

96.7

87.2

99.8

87.3

99.4

53.0

98.5

47.9

98.0

O

1

1a

O

2

2a

O

3a

3

O

4a

4

Cl

O

Cl 5

5a

O

6

6a

O

7 a

7a

(1) styrene, (2) 4-methylstyrene, (3) -methylstyrene, (4) trans--methylstyrene, (5) 3-chlorostyrene, (6) 1,2-dihydronaphthalene, (7) indene served as substrates. (1a) (S)-styrene oxide, (2a) 4-(S)-methyl-styrene oxide, (3a) (S)--methylstyrene oxide, (4a) (S)-trans--methylstyrene oxide, (5a) (S)-3-chlorostyrene oxide, (6a) (S)-1,2-dihydronaphthalene oxide, (7a) (1S,2R)-indene oxide were the corresponding epoxides synthesized biocatalytically.

b

390


References 1. Panke, S., Wubbolts, M.G., Schmid, A. and Witholt, B., Production of enantiopure styrene oxide by recombinant Escherichia coli synthesizing a two-component styrene monooxygenase. Biotechnol. Bioeng., 2000, 69, 91–100. 2. Park, J.B., Buehler, B., Habicher, T., Hauer, B., Panke, S., Witholt, B. and Schmid, A., The efficiency of recombinant Escherichia coli as biocatalyst for stereospecific epoxidation. Biotechnol. Bioeng., 2006, 95, 501–512. 3. Sambrook, J. and Russell, D.W., Molecular Cloning. A Laboratory Manual, 3rd edn, Nolan, C. (ed.). Cold Spring Harbor Laboratory Press: New York, 2001. 4. Wubbolts, M.G., Favre-Bulle, O. and Witholt, B., Biosynthesis of synthons in two-liquid-phase media. Biotechnol. Bioeng., 1996, 52, 301–308. 5. Schmid, A., Hofstetter, K., Feiten, H.J., Hollmann, F., and Witholt, B., Integrated biocatalytic synthesis on gram scale: the highly enantioselective preparation of chiral oxiranes with styrene monooxygenase. Adv. Synth. Catal., 2001, 343, 732–737.

12.9 Biotransformation of a-Bromo and a,a0 -Dibromo Alkanone

12.9

391

Biotransformation of a-Bromo and a, a 0 -Dibromo Alkanone into a-Hydroxyketone and a-Diketone by Spirulina platensis Takamitsu Utsukihara and C. Akira Horiuchi

-Hydroxy ketones are important as intermediates in organic synthesis.1,2 In a previous paper we found that a novel reaction of -bromo ketone under microwave irradiation gives the corresponding -hydroxy ketone in good yields.3 Biotransformation of -bromo and ,0 -dibromo alkanones was investigated with alga of Spirulina platensis. Biotransformation of -bromo ketone with S. platensis gave the corresponding -hydroxy ketone in good yields (80–95 %). It was found that ,0 -dibromo ketone is biocatalytically transformed into the -diketone and then is reduced into the -hydroxy ketone. In the case of 2,6-dibromo menthone, diosphenol (58 %), 1-hydroxy-3-methyl-6-isopropylcyclohexane-1,2-dione (15 %) and 2-hydroxy menthone (4 %) were obtained. This reaction affords a new, eco-friendly and convenient method for the synthesis of -hydroxy ketones. O

O Br

OH

Spirulina platensis 1:n = 0 2:n = 1 3:n = 2 4:n = 3

n 1– 4

n 1a – 4a

O

O Br

OH Spirulina platensis 5:R = H 6 : R = Me 7:R = F 8 : R = Cl 9 : R = Br

R

R

O

O Br

Br

O OH

Spirulina platensis

O +

n 2' – 3'

2' : n = 1 3' : n = 2

n

n

2a – 3a

Br

2b – 3b

OH Spirulina platensis

OH +

+

O

OH

O

O

O OH

Br

10

10a

10b

10c

Figure 12.8 Biotransformation of -bromo- and ,0 -dibromo alkanone by S. platensis?

392


12.9.1 12.9.1.1

Procedure 1: Cultivation of S. platensis Material and Equipment

• SOT medium:1 – NaHCO3 (16.8 g) – K2HPO4 (0.5 g) – NaNO3 (2.5 g) – K2SO4 (1 g) – NaCl (1 g) – MgSO47H2O (0.2 g) – CaCl22H2O (0.04 g) – FeSO47H2O (0.01 g) – Na2EDTA (0.08 g) – A5 solution (1 mL) – culture of S. platensis – distilled water 1000 mL • A5 solution: – H3BO3 (286 mg) – MnSO47H2O (250 mg) – ZnSO47H2O (22.2 mg) – CuSO45H2O (7.9 mg) – Na2MoO42H2O (2.1 mg) – distilled water 100 mL • • • •

filter paper one 200 mL Erlenmeyer flask fluorescent lamp air pump.

12.9.1.2

Procedure

1. SOT medium was prepared by mixing NaHCO3 (16.8 g), K2HPO4 (0.5 g), NaNO3 (2.5 g), K2SO4 (1 g), NaCl (1 g), MgSO47H2O (0.2 g), CaCl22H2O (0.04 g), FeSO47H2O (0.01 g), Na2EDTA (0.08 g) and A5 solution (1 mL) in distilled H2O (1000 mL). 2. A5 solution was H3BO3 (286 mg), MnSO47H2O (250 mg), ZnSO47H2O (22.2 mg), CuSO45H2O (7.9 mg) and Na2MoO42H2O (2.1 mg) dissolved in distilled H2O (100 mL). S. platensis was grown in SOT medium (pH 10–11) under continuous illumination provided by fluorescent lamps (2000 lx) with air bubbling at 25 C for 2 weeks. 3. The mixture was filtered to obtain the alga of S. platensis (yielded about 1 g L1 dry weight). 12.9.2

Procedure 2: Biotransformation of 2-Bromoacetophenone O

O Br



12.9.2.1 • • • • • • • • • • • •

393

Material and Equipment

S. platensis in SOT medium (100 mL) 2-bromoacetophenone (100 mg, 0.50 mmol) ethyl acetate ether hexane anhydrous sodium sulfate filter paper one 200 mL Erlenmeyer flask silica gel (Kieselgel 60 40–63 mm), 15 g fluorescent lamp shaker rotary evaporator.

12.9.2.2

Procedure

1. 2-Bromoacetophenone (100 mg, 0.50 mmol) was added to a suspended culture of S. platensis (adjusted pH 7.0, 1 g L1 as dry weight) in SOT medium (100 mL). The mixture was treated with a shaker (120 rpm) for 3 days at 25 C in the light (2000 lx). 2. At the end of the reaction, S. platensis was filtered off and the resulting mixture was extracted with EtOAc/Et2O (1:1). The organic layer was collected, dried over anhydrous sodium sulfate and concentrated using a rotary evaporator. The resulting oil was chromatographed on silica gel. Elution with hexane/ether (3:1) gave 2-hydroxyacetophenone (26 mg, 0.1 mmol). All the products were analysed by infrared (IR), 1H NMR and gas chromatography–mass spectrometry (GC–MS) analyses. 2-Hydroxyacetophenone. M.p. 85–86 C; 1H NMR (400 MHz, CDCl3): 3.54 (brs, 1H), 4.88 (s, 2H), 7.49 (t, 2H, J ¼ 7.7 Hz), 7.61 (t, 1H, J ¼ 7.4 Hz), 7.92 (d, 2H, J ¼ 7.6 Hz); 13C NMR (CDCl3): 65.4, 127.6, 128.9, 133.3, 134.2, 198.4. IR (KBr): 3428, 1687 cm1. MS (electron impact (EI)): m/z 136 (Mþ), 105, 77, 51. 12.9.3

Procedure 3: Biotransformation of 2,6-Dibromo Cyclohexanone O Br

O Br

Spirulina platensis

O OH

O +

12.9.3.1 • • • • • •


S. platensis in SOT medium (100 mL) 2,6-dibromo cyclohexanone (100 mg, 0.39 mmol) ethyl acetate ether hexane anhydrous sodium sulfate

394

• • • • • •


filter paper one 200 mL Erlenmeyer flask silica gel (Kieselgel 60 40–63 mm), 15 g fluorescent lamp shaker rotary evaporator.

12.9.3.2

Procedure

1. 2,6-Dibromo cyclohexanone (100 mg, 0.39 mmol) was added to suspended culture of S. platensis (adjusted pH 7.0, 1 g L1 as dry weight) in SOT medium (100 mL). The mixture was treated with a shaker (120 rpm) for 3 days at 25 C in the light (2000 lx). 2. At the end of the reaction, S. platensis was filtered off and the resulting mixture was extracted with EtOAc/Et2O (1:1). The organic layer was collected, dried over anhydrous sodium sulfate and concentrated using a rotary evaporator. 3. The resulting oil was chromatographed on silica gel. Elution with hexane/ether (3:1) gave 2-hydroxycyclohexanone (24 mg, 0.21 mmol) and 1,2-cyclohexanedione (0.8 mg, 0.007 mmol). All the products were analysed by IR, 1H NMR and GC–MS analyses. 2-Hydroxycyclohexanone. 1H NMR (400 MHz, CDCl3): 1.50–2.15 (m, 6H), 2.30– 2.50 (m, 2H), 3.66 (brs, 1H), 4.15 (ddd, 1H, J ¼ 1.6, 4.6, 8.8 Hz); 13C NMR (CDCl3): 23.4, 27.5, 36.7, 39.5, 75.3, 211.4. IR (neat): 3473, 1714 cm1. MS (EI): m/z 114 (Mþ), 96, 85, 70, 57, 44. 12.9.4

Procedure 4: Biotransformation of 2,6-Dibromo Menthone

Br


O

• • • • • • •


S. platensis in SOT medium (100 mL) 2,6-dibromo menthone (100 mg, 0.32 mmol) ethyl acetate ether hexane anhydrous sodium sulfate filter paper

OH +

+ O

Br

12.9.4.1

OH

O

O OH


• • • • •

395

one 200 mL Erlenmeyer flask silica gel (Kieselgel 60 40–63 mm), 15 g fluorescent lamp shaker rotary evaporator.

12.9.4.2

Procedure

1. 2,6-Dibromo menthone (100 mg, 0.32 mmol) was added to suspended culture of S. platensis (adjusted pH 7.0, 1 g L1 as dry weight) in SOT medium (100 mL). The mixture was treated with a shaker (120 rpm) for 3 days at 25 C in light (2000 lx). 2. At the end of the reaction, S. platensis was filtered off and the resulting mixture was extracted with EtOAc/Et2O (1:1). The organic layer was collected, dried over anhydrous sodium sulfate and concentrated using a rotary evaporator. 3. The resulting oil was chromatographed on silica gel. Elution with hexane/ether (3:1) gave diosphenol (15 mg, 0.09 mmol), 6-hydroxy-3-methyl-6-isopropylcyclohexane1,2-dione (3.7 mg, 0.02 mmol) and 2-hydroxy menthone (1.1 mg, 0.006 mmol). All the products were analysed by IR, 1H NMR and GC–MS. Diosphenol. 1H NMR (400 MHz, CDCl3): 1.00 (d, 6H), 1.12 (d, 3H), 2.36 (m, 1H), 6.34 (s, 1H); 13C NMR (CDCl3): 15.3, 19.5, 19.8, 22.3, 27.9, 30.9, 39.9, 138.1, 141.9, 197.4. IR (neat): 3425, 1670, 1620, 1160 cm1; MS (EI): 168 (Mþ), 153, 139, 126, 125, 108. 12.9.5

Conclusion

This is the first time that the biotransformation of -bromo and ,0 -dibromo ketone using S. platensis has been successfully accomplished. Although enantioselective -hydroxy ketones were not obtained, it was found that the hydroxylative biotransformation of -bromo and ,0 -dibromo alkanones using S. platensis affords a new synthetic method, which is more convenient, cleaner, and of lower energy than the chemical method used heretofore (see Tables 12.7 and 12.8).2–4 Biotransformation for -hydroxy ketone from -bromo ketone is no doubt attributable to the special properties of S. platensis system.

Table 12.7 Biotransformation of -bromo compounds by S. platensis Entry 1 2 3 4 5 6 7 8 9 a

Substrate

Day

Product (yield, %)a

1 2 3 4 5 6 7 8 9

1 1 1 5 3 3 3 3 3

1a (92) 2a (89) 3a (95) 4a (88) 5a (55) 6a (35) 7a (80) 8a (11) 9a (6)

Yield was determined by GC–MS peak area.

396

Whole-cell Oxidations and Dehalogenations Table 12.8 Biotransformation of ,0 -dibromo cycloalkanones by S. platensis Entry 1 2 3 a

Substrate

Day

Product (yield, %)a

20 30 10

3 5 3

2a (92) 2b (3) 3a (42) 3b (1) 10a (4) 10b (58) 10c (15)

Yield was determined by GC–MS peak area.

References 1. SOT: Spirulina–Ogawa–Terui. Ogawa, T. and Terui, G., Studies on the growth of Spindina platensis I. On the pure culture of Spindina platensis. J. Ferment. Technol., 1970, 48, 361. 2. Horiuchi, C.A., Takeda, A., Chai, W., Ohwada, K., Ji, S.-J. and Takahashi, T.T., A novel synthesis of -hydroxy- and ,0 -dihydroxyketone from -iodo and ,0 -diiodo ketone using photoirradiation. Tetrahedron Lett., 2003, 44, 9307. 3. Chai, W., Takeda, A., Hara, M., Ji, S.-J. and Horiuchi, C.A., Photo-irradiation of -halo carbonyl compounds: a novel synthesis of -hydroxy- and ,0 -dihydroxyketones. Tetrahedron, 2005, 61, 2453. 4. Utsukihara, T., Nakamura, H., Watanabe, M. and Horiuchi, C.A., Microwave-assisted synthesis of -hydroxy ketone and -diketone and pyrazine derivatives from -halo and ,0 -dibromo ketone. Tetrahedron Lett., 2006, 47, 9359.

Index

Note: Page references to figures are given in italic type; reference to tables are given in bold type. Abacavir 40–1 Acetylation 8 N-Acetyl-D-mannosamine (NAM) 33 N-Acetyl-D-neuraminic acid (NANA) 33 Acinetobacter calcoaceticus 332–5 Acremonium chrysogenum Acylation, 25–6, 36, 96, 367–8, Agrobacterium radiobacter 28 Alcalase 165–9 Alcohol dehydrogenases (ADH), see Ketoreductases 4–5, 48–52, 284–6 Alcohol reductases 8 Alcohols 288–90 esterification 36, 137–9 reduction 273–5 Aldehydes 271 Aldolases 52–4 Aldonic acids 323–4 Amano PS30 43 Amberlite XAD-1180 49 American Type Culture Collection 87 Amines dynamic kinetic resolution 148–52 free radical-mediated racemization 153–4 Amino acids 96–7, 314–17

Practical Methods for Biocatalysis and Biotransformations Ó 2009 John Wiley & Sons, Ltd

7-aminocephalosporic acid (7-ACA) 19–22, 65 Aminocyclitols 206–10 7-aminodesacetoxycephalosporanic acid (7-ADCA) 19, 22 Aminoshikimic acid 84 Aminotransferases 306–8 Amygdalin 242–3 Androgen receptor antagonists 37 Antibiotics 19–23 Aprepitant 51–2, 52 Arabidopsis thaliana 17 Arabinonucleosides 31 Archaea 90, 92, 101 Aspartate aminotransferase (AspAT) 306–8 Asymmetrization 35 Atorvastatin 28–9, 49–50 Azalactones, ethanolysis 162 Azides 232–4 1-azido disaccharides 232–5 Bacillus licheniformis 165 Bacillus sphaericus 314 Bacillus subtilis 190–7, 299 Bacillus subtilis protease 55–7 Bacteria 92, 112


398

Index

Baeyer-Villiger monooxygenase (BVMO) 299, 337–9 Baeyer-Villiger reactions 300, 301–4 Baker’s yeast 48 Basic Local Alignment Search Tool (BLAST) 89 Belgian Coordinated Collections of Microorganisms 87 Bioinformatics 88–90 Biotechnology 83–5 Biotransformation definition 3 BioWave reactor 362–5 Biphasic biocatalysis 59–61 Bisfuran alcohol 36 Brecanavir 36, 36–7 BRENDA 88 Buchner, Eduard 84 Candida antarctica lipase B (CALB) 24, 92, 133, 148, 170–2, 208–9 Candida rugosa lipase 110–11, 129–31 Carbamoylation 8 Carboxylic acid reductase 295–7 Cassette mutagenesis 107 CASTing 110 Centralbureau voor Schimmelcultures 87 Cephalexin 23 Cephalosporins 19–22 Chiral drug candidates 4–5 ChiroCLEC-PC 37 Chloroperoxidase (CPO) 327–9, 330–1 Cloning 91, 98–103 Clonostachys compactiuscula 25 Clopidogrel 43–4, 59–60, 291 Codons 96 Cofactors 86 see also Nicotinamide adenine dinucleotide (NADH) Combinatorial active site saturation test (CAST) 110 Complementary DNA (cDNA) 101 Corrin 69 Corynespora casiicola 376–8 Cosmids 99 Covalent enzyme attachment 62–3

Crispine A 319–21 Crosslinked enzyme aggregate (CLEA) 63, 266–7 Crosslinked enzyme crystals (CLEC) 63 Culture collections 87–8, 87, 93–4 Cultures, see Microbial cultures Cyanation 29 Cyanohydrin formation 255–8, 259–60, 266–7, 270–2 Cyclohexanone monooxygenase (CHMO) 332–5 Cyclopentadecanone monooxygenase (CPDMO) 344–9 CYP, see Cytochrome P450 Cytidine deaminase 39–40, 39 Cytochrome P450 9–11 microbial 12–13 Cytosine 96 Dealkylation 8 Deamination 8 Dean-Stark distillation 177 Dehydrated enzymes 56–7 Deoxyribonucleotide triphosphates (dNTP) 103 2-deoxyribose-5-phosphate aldolase (DERA) 52–3 Deracemization 320–1 Desymmetrization 41, 45–8, 125–8, 186–9, 341–3 Deutsche Sammlung von Mikroorganismen und Zellkulturen 87 Diasteroselectivity 30–4 Directed evolution 105–6, 106 DKR, see Dynamic kinetic resolution DNA 96–8 complementary (cDNA) 101 databases 89–90 noncoding 97–8 transcription 94–5 ligases 96, 100 polymerases 94–5, 103 sampling 90 shuffling 107–8 templates 98–103

Index

Drug metabolites 6–18 Dynamic kinetic resolution 141–164, 276–7

42–4, 137–9,

E-factor 64–5, 65–6 Enterobacter aerogenes 32 Entrapment (enzyme immobilization) 63–4 Environmental health and safety (EHS) 65 Environmental sampling 90–2 Enzyme activity databases 88–9 Enzyme induction 93 Epoxidation 8 Epoxide hydrolase 190–7 Error-prone PCR 106–7 Escherichia coli 27, 28 BL21(DE3) 291–2, 344–5 BVMO expression 337–9 carboxylic acid reductase 295–6 cytidine deaminase 39 enzyme induction 93 as expression host 111–12 JM101 385–9 Esterification 24–5, 25, 58, 160–4, 171–4 Eukaryotes 101 Eupergit C 63 ExPASy 90 Extremophiles 92, 93 Fagomine 212–17 Federal Drug Administration (FDA) 2 Fluvastatin 12, 29, 359–65 Fondaparinux 17 Free radicals 153–4 D-fructose-6-phosphate aldolase (FSA) 212–14 Fructose-1,6-bisphosphate aldolase (RAMA) 206–8 Fruit seed meal 236–9, 237, 269–71 Gene cloning 94–5, 102 testing 101–3 Gene identification, PCR 103–5 Gene synthesis 110 Generic drugs 1

399

Genetic code 96–8 Genetically modified microorganisms (GMMs) 5–6 Geotrichum candidum 48–9 Glucose isomerase 223–5 Glucose oxidase 323–5 Glucuronidation 246–9 Glycorandomization 18 Glycosidases 227 Glycosyl azides 232–4 Glycosylation 8, 16–18, 232–4 Glycosynthases 18, 227–9 Gordonae terrae 182–5 Green chemistry 63–6, 66 Halo-hydroxylation 327–9 Halohydrin dehydrogenase 199–200 Housekeeping genes 93 Humicola sp. lipase 125–7 Hydrolases 4, 35–6, 190–7, 341–2 see also Hydrolysis Hydrolysis 8, 23, 24, 48, 117–20, 135–6, 186–9, 339, 356–7, 359–65, 391–5 Hydroxylation 8, 9–10, 12, 206–8, 355–7, 359–65, 367–71 Hydroxynitrile lyase (HNL) 52–3, 255–7, 259–64 (S)-ibuprofen 157–61 Imidacloprid 355–8 Immobilization 61, 158 covalent attachment 62–3 noncovalent attachment 61–2 entrapment 63–4 hydroxynitrile lyase 266–7 T. versicolor laccase 243–4 Indels 109 Ionic liquids 39, 56 Irbesartan 9, 10 Isopropyl- -D-thiogalactopyranoside (IPTG) 93 Kazlaukas rules 46–7 Ketones 259–60, 278–82, 284–6 cyanohydrin synthesis 271 desymmetrization 125–8

400

Index

Ketones (Continued) reduction 288–90 see also Alcohol dehydrogenases (ADH) Ketoreductases 276–7, 278–82, 288–90, 289, 290 see also Alcohol dehydrogenases (ADH) Kinetic resolution 34–44, 117–20, 121–4, 129–31, 337–8 Laccases 15–16, 86–7 glycoside oxidation 240–4 -lactams 18–19 see also Cephalosporins Lactones 344–9 Lamivudine 39–40 LCA, see Life cycle analysis Leloir glycosyltransferases 17 Leptoxyphium fumago 327 Life cycle analysis (LCA) 65 Lipases 129–31, 134–6, 158–60, 173–80 Candida antarctica B 133, 148, 170–2, 208–9 immobilized 62 Pseudomonas fluorescens 41, 125 Lipolase 36 Liver cell microsomal fractions 11–12 horse 251–3 pig 245–9 Lobucavir 24–5, 25 Lotrafiban 38–9 Lovastatin 25, 47 Mandelic acid derivatives 43–4 Membrane reactors 64 Meso-trick 35 Metagenomics 90–2, 91 Microbacterium campoquemadoensis Microbial cultures 9, 111–12 collections 87–8, 87, 93–4 growth conditions 92–4 history 83–5 hosts 112–13 see also Culture collections Molecular biology 92–4 central dogma 94–5 enzyme tools 95–6

51

Molecular cloning 98 Monoamine oxidase 319–21 Monophasic biocatalysis 55–9 Monoterpenes 327–9 Montelukast 51, 52 Mortierella species 369–71 Motierella rammaniana 360–5 Mutagenesis 105–10 cassette mutagenesis 107 combinatorial methods 108–9 DNA shuffling 107–8 error-prone PCR 106–7 indels 109 neutral drift 109 rational enzyme design 109–10 Mutator strains 107 Mycophenolic acid 14, 14, 251–2 NADH 49, 86, 273–5 NAM 33 NANA aldolase 33 Napthalene 351–4 National Centre for Biotechnology Information (NCBI) 90 National Collection of Industrial Bacteria 87 Nelarabine 31–3, 31 Neutral drift 109 Nicotinamide adenine dinucleotide (NADH) 49, 86, 273–5 Nitriles 186–9 Noncoding DNA 97–8 Novozym 435 36, 37, 38, 45, 137–9 Nucleotide phosphorylases (NP) 30–2 Nucleotides 96 Odanacatib 42–3, 43 Olefins 355, 357 Oligosaccaride synthesis 227–9 Organic solvents 54–5 catalyst formulation 56–7 monophasic systems 55–6 solvent engineering 57–9 Origin of replication (ORI) 99

Index

Oxazolidines 173–4 Oxidation reactions 8, 11, 15–16, 299–304, 310–21, 323–6, 327–31, 333–4, 344–9, 351–4, 376–8, 385–9 P450, see Cytochrome P450 Palladium 148–52 Pasteur, Louis 83–4 PCR 103–5 Penicillin acylases 19 Penicillin G 19, 83–4, 84 pH memory effect 57 Phase I metabolic reactions 7, 8 Phase II metabolic reactions 8, 13–18 Phenylacetone monooxygenase (PAMO-P3) 299–303 Phenylalanine dehydrogenase (PheDH) 314–17 Photochemistry 299–304 Pig liver esterase (PLE) 93 -piperidine-2-carboxylate reductase (Pip2C) 310–12 Plantomycetes 117 Plasmids 98, 99 Polyesters 174–80, 179 Polymermatrices (as catalyst support) 63–4 Polymerase chain reaction (PCR) 103–5 error-prone 106–7 Posaconazole 45 Pregabalin 36 Prodrugs 23–4 Product lifetimes 1 ProSAR (protein sequence activity relationship) 6, 28–9 Proteases 121–4, 165 see also Bacillus subtilis protease Prunus dulcis 236–9 Prunus mume 269–72 Pseudomonas 21 Pseudomonas fluorescens lipase 41, 125 Pseudomonas mendocina 379–80 Pseudomonas putida 13, 310–11 PubMed 86

Racemization, see Dynamic kinetic resolution RAMA 206–8 Rational enzyme design 109–10 rDNA 84–5, 98 see also Cloning; DNA Reduction reactions 8 carboxylic acids 295–7 ketones 48–52, 284–6, 288–90 photochemical 303–4 Regioselectivity 18–29 Retro-claisenase 341–2 Reverse transcription 95 Rhamnulose-1-phosphate aldolase (rhAD) 203–5 Rhodococcus erythropolis NCIM 11540, 93, 186–8 Rhodococcus ruber 118–20 Ribavarin 24, 31 Riboflavin 84 RNA 96 Rosuvastatin 29 Roxifiban 43 Ruthenium 137–9 Saccharomyces cerevisiae, H402 x pTKL1 218–20 Sepabeads EC-EP 63 Sertraline 49 Shotgun libraries 100 Shuttle plasmids 98 Simvastatin 25–6 Solvents 39 biphasic systems 59–61 monophasic systems 55–7 organic 54–5 stereoselectivity and 59 see also Ionic liquids; Organic solvents; Supercritical fluids Spirulina platensis 391–5 Start codon 96 Statins 25, 28–9 Stavudine 27 Stenotrophomonas maltophilia 355

401

402

Index

Stop codon 96 Streptomyces griseoplanus 367–8 Streptomyces griseus 351–4 Streptomyces lividans 93 Streptomyces species 9, 21 Subtilisin Carlsberg 55–6, 165 Suicide vectors 99 Sulfatases 117 Sulfation 8 Supercritical fluids 56 T4MO 379–83 Taq polymerase 106–7 Terpenes 327–9 Thermoanaerobacter ethanolicus 284 Thermobifida fusca 299 Thermomyces lanuginosus 36 Thiol conjugation 8 Tissue preparations 11 see also Liver cell microsomal fractions TMPase 27 Trametes versicolor laccase 243–4 Transfection 98

Transformation 102 Tyrosinase 382–3 Urethane polyesters 174–6 Uridine diphosphate glucuronide transferase 14, 251–3 Uridine phosphorylase (URDP) 31 Valaciclovir 24 Vanillin 295–7 Vector promoters Vectors 98, 99 Viruses 98 Vitamin C 84

99

W110A secondary alcohol dehydrogenase 284–6 World Federation for Culture Collections 87 Yeasts

112

Zanamavir 33 Zeolite beta 133–36 Zidovudine 27

Practical Methods for Biocatalysis and Biotransformations

Practical Methods for Biocatalysis and Biotransformations 2

Practical Biotransformations: A Beginner's Guide

Practical Biotransformations: A Beginner's Guide

Biotransformations

Biotransformations

Biotransformations

Nanoscale Biocatalysis: Methods and Protocols (Methods in Molecular Biology 743)

Biotransformations and bioprocesses

Biotransformations and Bioprocesses

Microbial Enzymes and Biotransformations

Industrial Biotransformations

Biocatalysis: Fundamentals and Applications

Biocatalysis and Bioenergy

Biocatalysis and agricultural biotechnology

Biocatalysis and Biomolecular Engineering

Biocatalysis and Bioenergy

Biocatalysis: Fundamentals and Applications

Biocatalysis and Agricultural Biotechnology

Biocatalysis and Bioenergy

Industrial Biotransformations

Practical Fourier analysis for multigrid methods

Practical Fourier Analysis for Multigrid Methods

Practical Fibonacci Methods For Forex Trading

Enzyme Biocatalysis: Principles and Applications

Practical Methods for Biocatalysis and Biotransformations